Biomedical patches with aligned fibers

ABSTRACT

A three-dimensional electrospun nanofiber scaffold for use in repairing a defect in a tissue substrate is provided. The scaffold includes a flexible deposited fiber network of varying density including a first and second set of set of electrospun fibers. The second set of electrospun fibers is coupled to the first. A first portion of the flexible deposited fiber network includes a higher density of fibers than a second portion of the flexible deposited fiber network, and the tensile strength of first portion is higher than that of the second portion. The scaffold is sufficiently flexible to facilitate application of scaffold to uneven surfaces of the tissue substrate, and enables movement of the scaffold by the tissue substrate. The first and second set of fibers are configured to degrade within three months after application, and each fiber of the deposited fiber network has a diameter of 1-1000 nanometers.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application is a continuation of U.S. application Ser. No.15/497,691, filed Apr. 26, 2017, which is a continuation of U.S.application Ser. No. 13/703,210, now U.S. Pat. No. 10,149,749, filed onMar. 20, 2013, which is a national stage application under 35 U.S.C. §371 of International Application No. PCT/US2011/040691 filed on Jun. 16,2011, which claims the benefit of U.S. Provisional Application No.61/355,712, filed Jun. 17, 2010, all of which are incorporated herein byreference in their entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH & DEVELOPMENT

This invention was made with government support under Director's PioneerAward DP1 OD000798-04, awarded by the U.S. National Institutes ofHealth, and Award No. ECS-0335765, awarded by the U.S. National ScienceFoundation. The government has certain rights in the invention.

BACKGROUND

Numerous surgical procedures result in the perforation or removal ofbiological tissue, such as the water-tight fibrous membrane surroundingthe brain known as the dura mater. In some instances, such as minimallyinvasive neurosurgical procedures, relatively few small holes arecreated in the dura mater, while in others, such as the surgicalresection of advanced tumors, large sections of the dura mater may beremoved. In all of these cases, the tissue barrier surrounding the brainmust be repaired in order to prevent damage to cortical tissues andleakage of cerebrospinal fluid. To facilitate this repair, neurosurgeonsutilize sheets of polymeric materials or processed tissue that act likenative dura, known as dural substitutes.

At least some known dural substitutes utilized in neurosurgical clinicsare composed of an acellular collagen matrix obtained from isolatedbovine or porcine tissues. While generally accepted in the field, suchxenogenic dural substitutes may increase the incidence of adhesions andcontractures, transmit various zoonotic diseases to patients, andgenerally reduce patient outcome following surgery. Furthermore,processed collagenous grafts are exceedingly expensive, costing patientsand insurance companies thousands of dollars per procedure.

In addition while cell microarrays may be useful in biomedical researchand tissue engineering, at least some known techniques for producingsuch cell microarrays may be costly and time consuming, and may requirethe use of specialized, sophisticated instrumentation.

SUMMARY

One or more embodiments described herein provide structures having aplurality of aligned (e.g., radially aligned and/or polygonally aligned)fibers. When such a structure is used as a biomedical patch, thealignment of fibers as described herein may provide directional cuesthat influence cell propagation. For example, the structures providedmay promote new cell growth along the fibers, such that cell propagationin one or more desired directions may be achieved.

One or more structures provided may be created using an apparatus thatincludes one or more first electrodes that define an area and/orpartially circumscribe an area. For example, a single first electrodemay enclose the area, or a plurality of first electrode(s) may bepositioned on at least a portion of the perimeter of the area. A secondelectrode is positioned within the area. In exemplary embodiments, whenthe electrodes are electrically charged at a first polarity, and aspinneret dispensing a polymer (e.g., toward the second electrode) iselectrically charged at a second polarity opposite the first polarity,the dispensed polymer forms a plurality of fibers extending from thesecond electrode to the first electrodes. Further, electrodes withrounded (e.g., convex) surfaces may be arranged in an array, and afibrous structure created using such electrodes may include an array ofwells at positions corresponding to the positions of the electrodes.

In some embodiments, an artificial dura mater comprising at least ahydrophobic and biodegradable electrospun layer, wherein said layercomprises (a) at least one synthetic biomedical polymer and (b) fiberswith a diameter of 1-1000 nm is disclosed. In some embodiments, theartificial dura mater consists essentially of synthetic materials.

In some embodiments, a method of treating a subject having a defectivedura mater, the method comprising selecting an artificial dura materthat comprises at least one synthetic polymer and fibers with a diameterof 1-1000 nm, and applying said artificial dura mater proximate to saiddefective dura mater in said subject, is disclosed. In some embodiments,the artificial dura mater is as described elsewhere herein.

A three-dimensional electrospun nanofiber scaffold for use in repairinga defect in a tissue substrate is provided. The three-dimensionalelectrospun nanofiber scaffold includes a flexible deposited fibernetwork of varying density. The deposited fiber network further includesa first set of electrospun fibers including a first bioresorbablepolymer. The first bioresorbable polymer includes polyglycolic acid. Thedeposited fiber network further includes a second set of electrospunfibers including a second bioresorbable polymer. The second set offibers coupled is to the first set of fibers, and the firstbioresorbable polymer includes a different composition from the secondbioresorbable polymer. A first portion of the flexible deposited fibernetwork includes a higher density of fibers than a second portion of theflexible deposited fiber network, and the first portion has a highertensile strength than the second portion. The three-dimensionalelectrospun nanofiber scaffold is sufficiently flexible to facilitateapplication of the three-dimensional electrospun nanofiber scaffold touneven surfaces of the tissue substrate and to enable movement of thethree-dimensional electrospun nanofiber scaffold by the tissuesubstrate. The first set of fibers and the second set of fibers areconfigured to degrade within three months after application to thetissue substrate, and each fiber of the deposited fiber network has adiameter of 1-1000 nanometers.

A three-dimensional electrospun nanofiber scaffold for use in repairinga defect in a tissue substrate is provided. The three-dimensionalelectrospun nanofiber scaffold includes a flexible deposited fibernetwork of varying density. The deposited fiber network includes a firstset of electrospun fibers including a first bioresorbable polymer. Thefirst bioresorbable polymer includes glycolic acid. The deposited fibernetwork includes a second set of electrospun fibers including a secondbioresorbable polymer. The second set of fibers is coupled to the firstset of fibers and the first bioresorbable polymer has a differentcomposition from the second bioresorbable polymer. A first portion ofthe flexible deposited fiber network includes a higher density of fibersthan a second portion of the flexible deposited fiber network, and thefirst portion has a higher tensile strength than the second portion. Thethree-dimensional electrospun nanofiber scaffold is conformable to thedefect in the tissue substrate and is sufficiently flexible to enablemovement of the scaffold by the tissue substrate. The first set offibers and the second set of fibers are configured to separate from eachother within three months after application to the tissue substrate, andeach fiber of the deposited fiber network has a diameter of 1-1000nanometers.

A monolayer electrospun nanofiber patch for use in repairing a defect ina tissue substrate is provided. The monolayer electrospun nanofiberpatch includes a flexible deposited fiber network of varying density.The deposited fiber network includes a first set of electrospun fibersincluding a first bioresorbable polymer. The first bioresorbable polymerincludes glycolic acid. The monolayer electrospun nanofiber patchincludes a second set of electrospun fibers including a secondbioresorbable polymer. The second set of fibers is coupled to the firstset of fibers, and the first bioresorbable polymer includes a differentcomposition from the second bioresorbable polymer. An inner portion ofthe monolayer electrospun nanofiber patch includes a different fiberdensity than an outer portion of the monolayer electrospun nanofiberpatch. The monolayer electrospun nanofiber patch is conformable to thedefect in the tissue substrate, and is sufficiently flexible to enablemovement of the monolayer electrospun nanofiber patch by the tissuesubstrate. The first set of fibers and the second set of fibers areconfigured to degrade within three months after application to thetissue substrate, and each fiber of the deposited fiber network has adiameter of 1-1000 nanometers.

This summary introduces a subset of concepts that are described in moredetail below. This summary is not meant to identify essential features,and should not be read as limiting in any way the scope of the claimedsubject matter.

BRIEF DESCRIPTION OF THE DRAWINGS

The embodiments described herein may be better understood by referringto the following description in conjunction with the accompanyingdrawings.

FIG. 1 is a diagram illustrating a perspective view of an exampleelectrospinning system for producing a structure of radially alignedfibers.

FIG. 2 is a diagram illustrating an electric field generated by theelectrospinning system shown in FIG. 1.

FIG. 3 is a diagram of an electrode removed from the electrospinningsystem shown in FIG. 1 and having a plurality of fibers depositedthereon forming a biomedical patch.

FIG. 4 is a photograph of a biomedical patch including a plurality ofradially aligned electrospun fibers deposited on a peripheral electrode.

FIG. 5 is a scanning electron microscope (SEM) image of the biomedicalpatch shown in FIG. 4, further illustrating that the fibers of thebiomedical patch are radially aligned.

FIG. 6 is an illustration of a solid fiber spinneret.

FIG. 7 is an illustration of a hollow fiber spinneret.

FIG. 8 is an illustration of a biomedical patch layer with a pluralityof randomly oriented fibers, a biomedical patch layer with a pluralityof radially aligned fibers, and a multi-layer biomedical patch includingmultiple orders of fibers.

FIG. 9 is a diagram of a collector with a central electrode, an innerperipheral electrode defining an inner enclosed area, and an outerperipheral electrode defining an outer enclosed area.

FIG. 10 is a diagram of a concentric biomedical patch that may beproduced utilizing the collector shown in FIG. 9 in conjunction with theelectrospinning system shown in FIG. 1.

FIG. 11 is a flowchart of an exemplary method for producing a structureof radially aligned fibers using a peripheral electrode defining anenclosed area and a central electrode positioned approximately at acenter of the enclosed area.

FIG. 12 is a flowchart of an exemplary method for repairing a defect,insult, or void in a biological tissue.

FIG. 13 is a schematic illustration of a cellular infiltration of abiomedical patch from intact dural tissue apposing the edge of abiomedical patch.

FIG. 14A, FIG. 14B, FIG. 14C, and FIG. D are fluorescence micrographscomparing the migration of cells when dura tissues were cultured onscaffolds of radially aligned nanofibers and randomly orientednanofibers for 4 days. FIG. 14A is a fluorescence micrograph of duralfibroblasts stained with fluorescein diacetate (FDA) migrating alongradially aligned nanofibers. FIG. 14B is a fluorescence micrograph ofdural fibroblasts stained with FDA migrating along random fibers. FIG.14C is a fluorescence micrograph of dural fibroblasts stained with FDAmigrating along radially aligned nanofibers. FIG. 14D is a fluorescencemicrograph of dural fibroblasts stained with FDA migrating along randomfibers.

FIG. 15A, FIG. 15B, and FIG. 15C are schematic diagrams of a custom cellculture system designed to model the wound healing response of defectsor voids in a biological tissue. FIG. 15A is a diagram of a custom cellculture system including a metal ring. FIG. 15B is a diagram of a customcell culture system including a central silicone tube. FIG. 15C is a topview of a diagram of a custom cell culture system showing the locationof a central fiber scaffold and a surrounding region seeded withfibroblast cells.

FIG. 16A, FIG. 16B, FIG. 16C, and FIG. 16D are fluorescence micrographsshowing cell morphology and distribution on scaffolds of radiallyaligned nanofibers and randomly oriented nanofibers with and withoutfibronectin coating after incubation for 1 day. FIG. 16A is a micrographshowing cell morphology and distribution on scaffolds of radiallyaligned nanofibers. FIG. 16B is a micrograph showing cell morphology anddistribution on scaffolds of randomly aligned nanofibers. FIG. 16C is amicrograph showing cell morphology and distribution on scaffolds ofradially aligned nanofibers. FIG. 16D is a micrograph showing cellmorphology and distribution on scaffolds of randomly aligned nanofibers.

FIG. 17A, FIG. 17B, FIG. 17C, and FIG. 17D are fluorescence micrographsshowing the migration of dura fibroblasts seeded on fibronectin-coatedscaffolds of radially aligned nanofibers. FIG. 17A is a fluorescencemicrograph showing the migration of dura fibroblasts seeded onfibronectin-coated scaffolds of radially aligned nanofibers for 1 day.FIG. 17B is a fluorescence micrograph showing the migration of durafibroblasts seeded on fibronectin-coated scaffolds of radially alignednanofibers for 3 days. FIG. 17C is a fluorescence micrograph showing themigration of dura fibroblasts seeded on fibronectin-coated scaffolds ofradially aligned nanofibers for 7 days. FIG. 17D is a magnified view ofthe fluorescence micrograph of FIG. 17C showing the migration of durafibroblasts seeded on fibronectin-coated scaffolds of radially alignednanofibers for 7 days.

FIG. 18 is an illustration of a method utilized to determine the area ofremaining acellular region of the nanofiber scaffolds within thesimulated tissue defect.

FIG. 19 is a graph illustrating the acellular area remaining on thenanofiber scaffold within the simulated tissue defect as a function ofincubation time.

FIG. 20A, FIG. 20B, FIG. 20C, and FIG. 20D are fluorescence micrographsshowing live dural fibroblasts labeled with membrane dye on scaffolds ofradially aligned nanofibers with fibronectin coating. FIG. 20A is afluorescence micrographs showing live dural fibroblasts labeled withmembrane dye on scaffolds of radially aligned nanofibers withfibronectin coating after a 1-day culture. FIG. 20B is a fluorescencemicrographs showing live dural fibroblasts labeled with membrane dye onscaffolds of radially aligned nanofibers with fibronectin coating aftera 3-day culture. FIG. 20C is a fluorescence micrographs showing livedural fibroblasts labeled with membrane dye on scaffolds of radiallyaligned nanofibers with fibronectin coating after a 7-day culture. FIG.20D is a fluorescence micrographs showing live dural fibroblasts labeledwith membrane dye on scaffolds of radially aligned nanofibers withfibronectin coating after a 7-day culture and includes an inset of ahigh magnification image of the same.

FIG. 21A, FIG. 21B, FIG. 21C, and FIG. 21D are fluorescence micrographsdemonstrating the organization of cells and extracellular matrixadherent on scaffolds by immunostaining for type I collagen (green) andcell nuclei (blue). FIG. 21A is a fluorescence micrograph demonstratingthe organization of cells and extracellular matrix adherent on scaffoldsof radially aligned fibers by immunostaining for type I collagen (green)and cell nuclei (blue). FIG. 21B is a fluorescence micrographdemonstrating the organization of cells and extracellular matrixadherent on scaffolds of randomly oriented fibers by immunostaining fortype I collagen (green) and cell nuclei (blue). FIG. 21C is afluorescence micrograph demonstrating the organization of cells andextracellular matrix adherent on scaffolds of radially aligned fibers byimmunostaining for type I collagen (green) and cell nuclei (blue). FIG.21D is a fluorescence micrograph demonstrating the organization of cellsand extracellular matrix adherent on scaffolds of randomly orientedfibers by immunostaining for type I collagen (green) and cell nuclei(blue).

FIG. 22 is a graph illustrating the thickness of regenerated dura at thecenter of repaired dural defects over time.

FIG. 23 is a graph illustrating regenerative collagenous tissue contentover time.

FIG. 24 is a diagram illustrating a perspective view of an exampleelectrospinning system for producing a structure of fibers aligned inpolygons using an array of electrodes.

FIG. 25 is a diagram illustrating an elevation view of an examplemodular electrospinning collector.

FIG. 26 is a diagram illustrating an electric field generated by anelectrospinning system such as the electrospinning system shown in FIG.24.

FIG. 27A, FIG. 27B, FIG. 27C, FIG. 27D, FIG. 27E, and FIG. 27F aremicroscopy images of a membrane produced using a collector with an arrayof electrodes, such as the collector shown in FIG. 24. FIG. 27A is anoptical microscopy image of a membrane including an inset illustrating amagnification of the same. FIG. 27B is an optical microscopy image of amembrane including highlighted areas. FIG. 27C is a magnified opticalmicroscopy image of the highlighted area labeled 27C of FIG. 27B. FIG.27D is a magnified optical microscopy image of the highlighted arealabeled 27D of FIG. 27B. FIG. 27E is a magnified optical microscopyimage of the highlighted area labeled 27E of FIG. 27B. FIG. 27F is amagnified optical microscopy image of the highlighted area labeled 27Fof FIG. 27B.

FIG. 28A, FIG. 28B, FIG. 28C, and FIG. 28D are fluorescence microscopyimages illustrating cell growth in a membrane such as the membrane shownin FIGS. 27A-27F. FIG. 28A is an optical fluorescence microscopy imageof droplets containing cells placed within the wells of a fibermembrane. FIG. 28B is a fluorescence microscopy image array of cellsselectively adhered to the microwells within a nanofiber membrane. FIG.28C is a fluorescence microscopy image of seeded cell microarrays. FIG.28D is a fluorescence microscopy image of the same cell microarray shownin FIG. 28C after incubation for three days. 28A-28D are microscopyimages illustrating cell growth in a membrane such as the membrane shownin FIGS. 27A-27F.

FIG. 29A and FIG. 29B are microscopy images illustrating neuritepropagation in a membrane such as the membrane shown in FIGS. 27A-27F.FIG. 29A is an overlay of an optical microscopy image and a fluorescencemicroscopy image. FIG. 29B is an overlay of an optical microscopy imageand a fluorescence microscopy image adjacent to the region shown in FIG.29A.

FIG. 30A and FIG. 30B are overlays of an optical microscopy image and afluorescent microscopy image illustrating neuronal network formationfrom embryoid bodies in a membrane such as the membrane shown in FIGS.27A-27F. FIG. 30A is an overlay of an optical microscopy image and afluorescent microscopy image illustrating an embryoid body confinedwithin a microwell, while neurites extend peripherally along anunderlying fiber pattern. FIG. 30B is an overlay of an opticalmicroscopy image and a fluorescent microscopy image illustrating anembryoid body seeded on regions of uniaxially aligned nanofibers withina nanofiber array.

FIG. 31A, FIG. 31B, FIG. 31C, and FIG. 31D are scanning electronmicroscopy images illustrating membranes produced using a variety ofelectrode arrays. FIG. 31A is a scanning electron microscopy image of afiber membrane fabricated using a collector composed of hexagonal arraysof stainless steel beads. FIG. 31B is a scanning electron microscopyimage of a fiber membrane fabricated using a collector composed ofhexagonal arrays of stainless steel beads having a larger diameter thanthe stainless steel beads used to produce the membrane shown in FIG.31A. FIG. 31C is a scanning electron microscopy image of a fibermembrane fabricated using a collector composed of a close-packed squarearray of stainless steel beads. FIG. 31D is a scanning electronmicroscopy image of a fiber membrane produced using a collector composedof square arrays of stainless steel microbeads with a gradual increaseof the inter-electrode distance in one direction.

FIG. 32 is a diagram of a collector with peripheral electrodes partiallycircumscribing an area.

DETAILED DESCRIPTION

Embodiments provided herein facilitate repairing biological tissue withthe use of a biomedical patch including a plurality of fibers. Suchfibers may have a very small cross-sectional diameter (e.g., from 1-1000nanometers) and, accordingly, may be referred to as nanofibers. Whilebiomedical patches are described herein with reference to dura mater anduse as a dural substitute, embodiments described may be applied to anybiological tissue. Moreover, although described as biomedical patches,structures with aligned fibers may be used for other purposes.Accordingly, embodiments described are not limited to biomedicalpatches.

In operation, biomedical patches provided herein facilitate cell growthand may be referred to as “membranes,” “scaffolds,” “matrices,” or“substrates.” Such biomedical patches further facilitate cell migrationfrom a perimeter of the patch to a center of the biomedical patch.Biomedical patches with aligned fibers, as described herein, may promotesignificantly faster healing and/or regeneration of tissue such as thedura mater than substitutes lacking nanoscopic organization anddirectional cues.

Dura mater is a membranous connective tissue located at the outermost ofthe three layers of the meninges surrounding the brain and spinal cord,which covers and supports the dural sinuses and carries blood from thebrain towards the heart. Dural substitutes are often needed after aneurosurgical procedure to repair, expand, or replace the incised,damaged, or resected dura mater.

Although many efforts have been made, the challenge to develop asuitable dural substitute has been met with limited success. Autografts(e.g., fascia lata, temporalis fascia, and pericranium) are preferablebecause they do not provoke severe inflammatory or immunologicreactions. Potential drawbacks of autografts include the difficulty inachieving a watertight closure, formation of scar tissue, insufficientlyaccessible graft materials to close large dural defects, increased riskof infection, donor site morbidity, and the need for an additionaloperative site. Allografts and xenografts are often associated withadverse effects such as graft dissolution, encapsulation, foreign bodyreaction, scarring, adhesion formation, and toxicity-induced sideeffects from immunosuppressive regimens. Lyophilized human dura mater asa dural substitute has also been reported as a source of transmittablediseases, specifically involving prions, such as Creutzfeldt-Jakobdisease.

In terms of materials, non-absorbable synthetic polymers, such assilicone and expanded polytetrafluoroethylene (ePTFE), often causeserious complications that may include induction of granulation tissueformation due to their chronic stimulation of the foreign body response.Natural absorbable polymers, including collagen, fibrin, and cellulose,may present a risk of infection and disease transmission. As a result,synthetic polymers such as poly(3-hydroxybutyrate-co-3-hydroxyvalerate)(PHBV), poly(lactic acid) (PLA), polyglycolic acid (PGA), PLA-PCL-PGAternary copolymers, and hydroxyethylmethacrylate hydrogels have recentlyattracted attention as biodegradable implant materials for dural repair.Methods and systems described herein may be practiced with thesematerials and/or any biomedical polymer.

In order to facilitate successful regeneration and/or repair of the duramater following surgery, a synthetic dural substitute or biomedicalpatch should promote: i) adhesion of dural fibroblasts (the primary celltype present in the dura) to the surface of the biomedical patch; ii)migration of dural fibroblasts from the periphery of the biomedicalpatch toward the center; and iii) minimal immune response. To date,synthetic dural substitutes have been tested only in the form of foils,films, meshes, glues, and hydrogels. Due to the isotropic surfaceproperties, such substitutes are not well-suited for cell attachment anddirected, inward migration.

This problem can be potentially solved by fabricating the polymers asnanoscale fibers with a specific order and organization. For example,the speed of cell migration may be very low on flat, isotropic surfaces,whereas cells may migrate over a very long distance in a highlycorrelated fashion with constant velocity on a uniaxially aligned,fibrous scaffold.

Electrospinning is an enabling technique which can produce nanoscalefibers from a large number of polymers. The electrospun nanofibers aretypically collected as a randomly-oriented, nonwoven mat. Uniaxiallyaligned arrays of nanofibers can also be obtained under certainconditions, specifically when employing an air-gap collector or amandrel rotating at a high speed. However, uniaxially aligned nanofiberscaffolds promote cell migration only along one specific direction andare thus not ideally suited as dural substitutes.

In order to promote cell migration from the surrounding tissue to thecenter of a dural defect and shorten the time for healing andregeneration of dura mater, a surface patterned with aligned (e.g.,aligned radially and/or in one or more polygons), nanoscale featureswould be highly advantageous as an artificial dural substitute. Morespecifically, scaffolds constructed with aligned nanofibers could meetsuch a demand by guiding and enhancing cell migration from the edge of adural defect to the center.

Many polymers are available for use in electrospinning. In someembodiments described herein, nanofibers for dura substitutes areproduced as the electrospun polymer from poly(ε-caprolactone) (PCL), anFDA approved, semicrystalline polyester that can degrade via hydrolysisof its ester linkages under physiological conditions with nontoxicdegradation products. This polymer has been extensively utilized andstudied in the human body as a material for fabrication of drug deliverycarriers, sutures, or adhesion barriers. As described herein,electrospun PCL nanofibers may be aligned to generate scaffolds that areuseful as dural substitutes.

Embodiments provided herein facilitate producing a novel type ofartificial tissue substitute including a polymeric nanofiber material,which is formed through a novel method of electrospinning. Thispolymeric material includes non-woven nanofibers (e.g., fibers having adiameter of 1-1000 nanometers) which are aligned within a materialsheet.

In exemplary embodiments, a material with aligned nanofibers is formedthrough a novel method of electrospinning that employs a collectorincluding one or more first, or “peripheral,” electrodes defining anarea and/or at least partially circumscribing the area, and a second, or“inner,” electrode positioned within the area. When the electrodes areelectrically charged at a first polarity, and a spinneret dispensing apolymer (e.g., toward the inner electrode) is electrically charged at asecond polarity opposite the first polarity, the dispensed polymer formsa plurality of fibers extending from the inner electrode to theperipheral electrode(s). Electrodes may include a rounded (e.g., convex)surface, such that a depression, or “well”, is formed in theelectrode-facing side of a structure of fibers. Alternatively,electrodes may include a concave surface, such that a well is formed inthe side of the structure facing away from the electrodes.

In some embodiments, the collector includes a single inner electrode anda single peripheral electrode. In other embodiments, the collectorincludes a plurality of peripheral electrodes, and the dispensed polymermay form fibers extending between such peripheral electrodes in additionto fibers extending from the inner electrode to one or more of theperipheral electrodes.

Further, in some embodiments, multiple areas are defined and/orpartially circumscribed by peripheral electrodes. For example, an innerperipheral electrode may define an inner enclosed area surrounding theinner electrode, and an outer peripheral electrode may define an outerenclosed area surrounding the inner peripheral electrode. In otherembodiments, electrodes are arranged in an array, such as a grid and/orother polygonal pattern (e.g., a hexagonal pattern), and multiple,partially overlapping areas may be defined by such electrodes. Forexample, an inner electrode of one area may function as a peripheralelectrode of another area. In such embodiments, the dispensed polymermay form fibers extending between the electrodes of the collector, suchthat the fibers define the sides of a plurality of polygons, with theelectrodes positioned at the vertices of the polygons.

Unlike known nanofiber structures, aligned nanofiber materials providedherein are capable of presenting nanoscale topographical cues to localcells that enhance and direct cell migration (e.g., throughout thematerial sheet or into the center of the material sheet). As a result,aligned nanofiber materials may induce faster cellular migration andpopulation than randomly oriented materials, such as processedgold-standard collagen matrices. Materials described herein may beparticularly useful as a substrate for various types of biomedicalpatches or grafts designed to induce wound protection, closure, healing,repair, and/or tissue regeneration.

A scaffold of aligned nanofibers, as described herein, possessessignificant potential as an artificial dural substitute, in that it iscapable of encouraging robust cell migration from apposed intact duraand promoting rapid cellular population of the nanofiber matrix requiredto induce dural repair. In addition, such nanofiber materials offer theadvantage of being inexpensive to produce, fully customizable, andresorbable. Nanofiber dural substitutes may also reduce the risk ofcontractures and fully eliminate the risk of transmitted zoonoticdisease when applied intraoperatively, generally improving patientoutcomes following surgery.

Inner Electrode and Peripheral Electrode(s)

FIG. 1 is a diagram illustrating a perspective view of an exemplaryelectrospinning system 100 for producing a structure of radially alignedfibers. System 100 includes a collector 105 with a first electrode 110,which may be referred to as a peripheral electrode, and a secondelectrode 115, which may be referred to as an inner electrode or centralelectrode. System 100 also includes a spinneret 120. Peripheralelectrode 110 defines an enclosed area 125, and central electrode 115 ispositioned approximately at a center of enclosed area 125.

System 100 is configured to create an electric potential betweencollector 105 and spinneret 120. In one embodiment, peripheral electrode110 and central electrode 115 are configured to be electrically chargedat a first amplitude and/or polarity. For example, peripheral electrode110 and central electrode 115 may be electrically coupled to a powersupply 130 via a conductor 135. Power supply 130 is configured to chargeperipheral electrode 110 and central electrode 115 at the firstamplitude and/or polarity via conductor 135.

In the embodiment illustrated in FIG. 1, peripheral electrode 110 is aring defining an enclosed area 125 which is circular. For example,circular enclosed area 125 may have a diameter of between 1 centimeterand 20 centimeters. In other embodiments, peripheral electrode 110 maybe any shape suitable for use with the methods described herein. Forexample, peripheral electrode 110 may define an elliptical, ovular,rectangular, square, triangular, and/or other rectilinear or curvilinearenclosed area 125. In some embodiments, peripheral electrode 110 definesan enclosed area 125 of between 5 square centimeters and 100 squarecentimeters. Peripheral electrode 110 may have a height 112 of between0.5 and 2.0 centimeters. Central electrode 115 may include a metallicneedle and/or any other structure terminating in a point or set ofpoints.

In one embodiment, enclosed area 125 defines a horizontal plane 127.Spinneret 120 is aligned with central electrode 115 and verticallyoffset from horizontal plane 127 at a variable distance. For example,spinneret 120 may be vertically offset from horizontal plane 127 at adistance of 1 centimeter to 100 centimeters.

Spinneret 120 is configured to dispense a polymer 140 while electricallycharged at a second amplitude and/or polarity opposite the firstpolarity. As shown in FIG. 1, spinneret 120 is electrically coupled topower supply 130 by a conductor 145. Power supply 130 is configured tocharge spinneret 120 at the second amplitude and/or polarity viaconductor 145. In some embodiments, power supply 130 provides a directcurrent (DC) voltage (e.g., between 10 kilovolts and 17 kilovolts). Inone embodiment, conductor 145 is charged positively, and conductor 135is charged negatively or grounded. In some embodiments, power supply 130is configured to allow adjustment of a current, a voltage, and/or apower.

In one embodiment, spinneret 120 is coupled to a syringe 150 containingpolymer 140 in a liquid solution form. Syringe 150 may be operatedmanually or by a syringe pump 155. In an exemplary embodiment, spinneret120 is a metallic needle having an aperture between 100 micrometers and2 millimeters in diameter.

As syringe 150 pressurizes polymer 140, spinneret 120 dispenses polymer140 as a stream 160. Stream 160 has a diameter approximately equal tothe aperture diameter of spinneret 120. Stream 160 descends towardcollector 105. For example, stream 160 may fall downward under theinfluence of gravity and/or may be attracted downward by a chargedconductive surface 162 positioned below collector 105. For example,conductive surface 162 may be electrically coupled to conductor 135 andcharged at the same amplitude and/or polarity as peripheral electrode110 and central electrode 115. As stream 160 descends, polymer 140 formsone or more solid polymeric fibers 165.

In some embodiments, a mask 164 composed of a conducting ornon-conducting material is applied to collector 105 to manipulatedeposition of fibers 165. For example, mask 164 may be positionedbetween spinneret 120 and collector 105 such that no fibers 165 aredeposited on collector 105 beneath mask 164. Moreover, mask 164 may beused as a time-variant mask by adjusting its position while spinneret120 dispenses polymer 140, facilitating spatial variation of fiberdensity on collector 105. While mask 164 is shown as circular, mask 164may have any shape (e.g., rectangular or semi-circular) and sizesuitable for use with system 100. Alternatively, or in addition,deposition of fibers 165 on collector 105 may be manipulated byadjusting the position of collector 105 with respect to spinneret 120 orby spatially varying the electrical potential applied between thespinneret 120 and/or the electrodes making up the collector 105. Forexample, positioning one side of collector 105 directly beneathspinneret 120 may cause more fibers 165 to be deposited on that sidethan are deposited on the opposite side of collector 105.

FIG. 2 is a diagram 200 illustrating an electric field generated bysystem 100. Diagram 200 shows a two dimensional, cross-sectional view ofelectric field strength vectors between spinneret 120 and peripheralelectrode 110 and central electrode 115 of collector 105 (shown in FIG.1). Unlike known electrospinning systems, the electric field vectors(stream lines) in the vicinity of the collector are split into twopopulations, pointing toward the peripheral electrode 110 and pointingtoward the central electrode 115.

Neglecting the effect of charges on the polymeric fibers, the electricalpotential field can be calculated using the Poisson equation,

${{\nabla^{2}V} = \frac{- \rho}{ɛ}},$where V is the electrical potential, ε is the electrical permittivity ofair, and ρ is the spatial charge density. The electrical field, E, canthen be calculated by taking the negative gradient of the electricalpotential field, E=−∇V. Here, the electrical field was calculated toverify the alignment effect demonstrated by deposited fibers, which wasperformed using the software COMSOL3.3.

FIG. 3 is a diagram of peripheral electrode 110 removed fromelectrospinning system 100 (shown in FIG. 1) and having a plurality offibers 165 deposited thereon forming a biomedical patch 170. Fibers 165extend radially between a center 175 corresponding to the position ofcentral electrode 115 (shown in FIG. 1) and a perimeter 178corresponding to the position of peripheral electrode 110. For example,perimeter 178 may be a circular perimeter about center 175 defining adiameter of between 1 centimeter and 6 centimeters.

Biomedical patch 170 is illustrated with a small quantity of fibers 165in FIG. 3 for clarity. In some embodiments, biomedical patch 170includes thousands, tens of thousands, hundreds of thousands, or morefibers 165, evenly distributed throughout enclosed area 125 (shown inFIG. 1) of peripheral electrode 110. Even with millions of fibers 165,biomedical patch 170 is flexible and/or pliable, facilitatingapplication of biomedical patch 170 to uneven biological tissuesurfaces, such as the surface of the dura mater.

The radial alignment of fibers 165 demonstrates the shortest possiblepath between perimeter 178 and center 175. Accordingly, biomedical patch170 also facilitates cell migration directly from perimeter 178 tocenter 175, enabling a reduction in time required for cells toinfiltrate and populate applied biomedical patch, and for native tissueto regenerate.

Fibers 165 have a diameter of 1-1000 nanometers. In one embodiment,fibers have a diameter of approximately 220 nanometers (e.g., 215 nm to225 nm). The diameter of the fibers 165, thickness of the biomedicalpatch 170, and/or fiber density within the biomedical patch 170 mayaffect the durability (e.g., tensile strength) of biomedical patch 170.Biomedical patch 170 may be produced with various mechanical propertiesby varying the thickness and/or the fiber density of the biomedicalpatch 170 by operating electrospinning system 100 for relatively longeror shorter durations.

FIG. 4 is a photograph 300 of a biomedical patch 305 including aplurality of radially aligned electrospun fibers deposited on aperipheral electrode 110. FIG. 5 is a scanning electron microscope (SEM)image 310 of biomedical patch 305, further illustrating that the fibersof biomedical patch 305 are radially aligned.

Referring to FIGS. 1 and 3, fibers 165 may be solid or hollow. In someembodiments, the size and/or structure of fibers 165 is determined bythe design of spinneret 120. FIG. 6 is an illustration of a solid fiberspinneret 120A. Solid fiber spinneret 120A includes a conical body 180defining a center line 182. At a dispensing end 184, conical body 180includes an annulus 186. Annulus 186 defines a circular aperture 190A,through which polymer 140 may be dispensed. Fibers 165 produced withsolid fiber spinneret 120A have a solid composition.

FIG. 7 is an illustration of a hollow fiber spinneret 120B. Like solidfiber spinneret 120A, hollow fiber spinneret 120B includes a conicalbody 180 with an annulus 186 at a dispensing end 184. Hollow fiberspinneret 120B also includes a central body 188B positioned withinannulus 186. Annulus 186 and central body 188B define an annularaperture 190B. Accordingly, when polymer 140 is dispensed by hollowfiber spinneret 120B, fibers 165 have a hollow composition, with anexterior wall surrounding a cavity. The exterior wall of a fiber 165dispensed by hollow fiber spinneret 120B defines an outer diametercorresponding to the inner diameter of annulus 186 and an inner diametercorresponding to the diameter of central body 188B. Accordingly, theouter diameter and inner diameter of hollow fibers 165 may be adjustedby adjusting the diameters of annulus 186 and central body 188B.

Hollow fiber spinneret 120B facilitates incorporating a substance, suchas a biological agent, growth factor, and/or a drug (e.g., achemotherapeutic substance), into biomedical patch 170. For example, thesubstance may be deposited within a cavity defined by hollow fibers 165of biomedical patch 170. In one embodiment, polymer 140 is selected tocreate porous and/or semi-soluble fibers 165, and the substance isdispensed from the cavity through fibers 165. In another embodiment,polymer 140 is degradable, and the substance is dispensed as fibers 165degrade in vivo. For example, fibers 165 may be configured to degradewithin 12 months, 6 months, or 3 months. The degradation rate of polymer140 may be manipulated by adjusting a ratio of constituent polymerswithin polymer 140.

In another embodiment, a substance is delivered by solid fibers 165. Forexample, a solid fiber 165 may be created from a polymer 140 includingthe substance in solution. As solid fiber 165 degrades, the substance isreleased into the surrounding tissue.

As shown in FIGS. 6 and 7, annulus 186 is perpendicular to center line182. In an alternative embodiment, annulus 186 is oblique (e.g.,oriented at an acute or obtuse angle) with respect to center line 182.The outside diameter of fibers 165 may be determined by the insidediameter of annulus 186.

Some embodiments facilitate producing a biomedical patch having radiallyaligned fibers and non-radially aligned fibers. For example, radiallyaligned fibers may be deposited into a first layer, and non-radiallyaligned fibers may be deposited into a second layer. Alternatively,radially aligned non-radially aligned fibers may be deposited into asingle layer (e.g., simultaneously, sequentially, and/or alternately).Referring to FIG. 1, system 100 may be used to create randomly orientedfibers by charging or grounding conductive surface 162. Optionally,peripheral electrode 110 and central electrode 115 may be uncharged orungrounded (e.g., decoupled from conductor 135).

FIG. 8 is an illustration of a biomedical patch layer 400 with aplurality of randomly oriented fibers 405 and a biomedical patch layer410 with a plurality of radially aligned fibers 415. As shown in FIG. 8,biomedical patch layers 400 and 410 may be combined (e.g., overlaid) toproduce a multi-layer biomedical patch 420 with both randomly orientedfibers 405 and radially aligned fibers 415, or any other combination ofany number or type of fiber layers. Combining non-radially alignedfibers 405 and radially aligned fibers 415 facilitates providing abiomedical patch that promotes cell migration to a center of thebiomedical patch while exhibiting potentially greater durability (e.g.,tensile strength) than a biomedical patch having only radially alignedfibers 415. Combining non-radially aligned fibers 405 and radiallyaligned fibers 415 may also enable spatial control of cell migration andinfiltration along an axis perpendicular to the plane of the biomedicalpatch, facilitating the formation and organization of specific layers ofcells and/or extracellular matrix proteins resembling natural tissuestrata.

In some embodiments, multiple biomedical patch layers 410 with radiallyaligned fibers 415 may be combined to create a multi-layer biomedicalpatch. For example, referring to FIGS. 1 and 3, after depositing a firstset of fibers on collector 105, one may wait for the first set of fibers165 to solidify completely or cure and then deposit a second set offibers 165 on collector 105. The second set of fibers 165 may bedeposited directly over the first set of fibers 165 on collector 105.Alternatively, the first set of fibers 165 may be removed from collector105, and the second set of fibers 165 may be deposited on conductivesurface 162 and/or collector 105 and then removed and overlaid on thefirst set of fibers 165. Such embodiments facilitate increaseddurability of the biomedical patch, and added spatial control of cellmigration/activity, even where only radially aligned fibers are used. Insome embodiments, a hydrogel or polymeric scaffold may be disposedbetween biomedical patch layers 400 and/or biomedical patch layers 410.

A multi-layered biomedical patch may be useful for dural grafts as wellas other tissue engineering applications. Sequential layers of fiberscan be created with varying orders (e.g., radially aligned or randomlyoriented) and densities (e.g., low or high fiber density), which mayallow specific types of cells to infiltrate and populate select layersof the artificial biomedical patch. For example, biomedical patchescontaining a high fiber density generally prohibit cellular migrationand infiltration, while biomedical patches containing a low fiberdensity generally enhance cellular migration and infiltration.

Overall, the ability to form multi-layered fiber materials, as describedherein, may be extremely beneficial in the construction of biomedicalpatches designed to recapitulate the natural multi-laminar structure ofnot only dura mater, but also other biological tissues such as skin,heart valve leaflets, pericardium, and/or any other biological tissue.Furthermore, one or more layers of a biomedical patch may be fabricatedfrom biodegradable polymers such that the resulting nanofiber materialsfully resorb following implantation. Manipulation of the chemicalcomposition of the polymers utilized to fabricate these scaffolds mayfurther allow for specific control of the rate of degradation and/orresorption of a biomedical patch following implantation.

Some embodiments provide a biomedical patch including a plurality ofnested (e.g., concentric) areas. FIG. 9 is a diagram of a collector 505with a central electrode 115, a first or inner peripheral electrode 510defining a first or inner enclosed area 515, and a second or outerperipheral electrode 520 defining a second or outer enclosed area 525that is larger than the inner enclosed area 515. In some embodiments,outer peripheral electrode 520 is concentrically oriented with innerperipheral electrode 510. While inner peripheral electrode 510 and outerperipheral electrode 520 are shown as defining circular enclosed areas515, 525 in FIG. 9, inner peripheral electrode 510 and outer peripheralelectrode 520 may define enclosed areas 515, 525 of any shape suitablefor use with the methods described herein. Moreover, inner enclosed area515 and outer enclosed area 525 may have different shapes and/ordifferent centers.

In operation with electrospinning system 100 (shown in FIG. 1), centralelectrode 115 and inner peripheral collector 505 are charged at thefirst amplitude and/or polarity (opposite the polarity at whichspinneret 120 is charged) while spinneret 120 dispenses polymer 140 asstream 160. Stream 160 descends toward collector 505 and forms one ormore fibers 530 extending from central electrode 115 to inner peripheralelectrode 510.

The charge of the first polarity is removed from central electrode 115(e.g., by decoupling central electrode 115 from conductor 135), andouter peripheral electrode 520 is charged at the first amplitude and/orpolarity. Spinneret 120 dispenses polymer 140 as stream 160, whichdescends toward collector 505 and forms one or more fibers 535 extendingfrom inner peripheral electrode 510 to outer peripheral electrode 520.Together, fibers 530 and 535 form a concentric biomedical patch 550, asshown in FIG. 10. In some embodiments, the charge is not removed fromcentral electrode 115 prior to depositing fibers 535 between innerperipheral electrode 510 and outer peripheral electrode 520.

FIG. 10 is a diagram of a concentric biomedical patch 550 that may beproduced with collector 505 (shown in FIG. 9). Fibers 530 define aninner area 555, shown as a circle extending from a center 560 to aninner perimeter 565. An outer area 570 includes fibers 535 extendingapproximately from inner perimeter 565 (e.g., about 100 μm to 2000 μminside inner perimeter 565) to an outer perimeter 575. Fibers 535 areoriented radially or approximately (e.g., within 1, 3, or 5 degrees)radially with respect to center 560.

As shown in FIG. 10, inner area 555 and outer area 570 may overlap in anoverlapping area 580. In one embodiment, overlapping area 580corresponds to a thickness of inner peripheral ring 510 (shown in FIG.8). Similar to FIG. 3, concentric biomedical patch 550 is shown in FIG.10 with a small quantity of fibers 530 and 535 for clarity. In someembodiments, inner area 555 and outer area 570 each include thousands,tens of thousands, hundreds of thousands, or more fibers 530 and 535,respectively. Fibers 530 and fibers 535 may be coupled to each other inoverlapping area 580. For example, fibers 535 may be deposited beforefibers 530 have completely solidified (or vice versa). In someembodiments, fibers 530 and fibers 535 are deposited on collector 505(shown in FIG. 9) simultaneously or in an alternating manner.

Embodiments such as those shown in FIGS. 9 and 10 facilitate providing abiomedical patch having a relatively consistent fiber densitythroughout. For contrast, if fibers 530 extended from center 560 toouter perimeter 575, the fiber density at center 560 would beconsiderably higher than the fiber density at outer perimeter 575. Lowperipheral fiber density may compromise durability of a biomedical patchnear an outer perimeter, especially at large diameters (e.g., above 5 or6 centimeters). Accordingly, such embodiments further facilitateproviding a biomedical patch of large diameter (e.g., up to 10 or 12centimeters) while maintaining durability of the biomedical patch. Insome embodiments, a layer of non-radially aligned fibers is combinedwith biomedical patch 550, as described above with regard to FIG. 8,which may further enhance durability of biomedical patch 550.

In some embodiments, the spatial fiber density within inner area 555 isdifferent from the spatial fiber density within outer area 570. In oneexample, fibers 530 are deposited between central electrode 115 andinner peripheral electrode 510 for a first duration, and fibers 535 aredeposited between inner peripheral electrode 510 and outer peripheralelectrode 520 for a second duration.

While collector 505 and concentric biomedical patch 550 are illustratedwith circular inner and outer areas, any quantity and shape ofperipheral electrodes may be used to create any number of distinct fiberareas within a biomedical patch.

FIG. 11 is a flowchart of an exemplary method 600 for producing astructure of radially aligned fibers using a peripheral electrodedefining an enclosed area and a central electrode positionedapproximately at a center of the enclosed area. While one embodiment ofmethod 600 is shown in FIG. 11, it is contemplated that any of theoperations illustrated may be omitted and that the operations may beperformed in a different order than is shown.

Method 600 includes electrically charging 605 the peripheral electrodeand the central electrode at a first amplitude and/or polarity (e.g.,negatively charging or grounding). A spinneret approximately alignedwith the central electrode is electrically charged 610 at a secondamplitude and/or polarity opposite the first amplitude and/or polarity(e.g., positively charged).

A polymer (e.g., a liquid polymer) is dispensed 615 from the spinneret.In an exemplary embodiment, dispensing 615 the polymer forms a pluralityof polymeric fibers extending from the central electrode to theperipheral electrode to create a layer of radially aligned fibers.

Some embodiments facilitate creating a concentric structure of radiallyaligned fibers using multiple peripheral electrodes. In one embodiment,the peripheral electrode is an inner peripheral electrode. An outerperipheral electrode defining an outer enclosed area larger than theinner enclosed area is electrically charged 620 at the first amplitudeand/or polarity. The electrical charge may or may not be removed 622from the central electrode and/or the inner peripheral electrode. Thepolymer is dispensed 625 from the spinneret to create an outer area ofradially aligned fibers extending from the inner peripheral electrode tothe outer peripheral electrode.

Furthermore, some embodiments facilitate creating a multi-layeredstructure including both radially aligned fibers and non-radiallyaligned fibers. The electrical charge is removed 630 from the peripheralelectrode(s) and the central electrode. A conductive surface below thelayer of radially aligned fibers is electrically charged 635 at thefirst amplitude and/or polarity. The polymer is dispensed 640 from thespinneret to create a layer of non-radially aligned (e.g., randomlyoriented and/or uniaxially aligned) fibers over the layer of radiallyaligned fibers.

FIG. 12 is a flowchart of an exemplary method 700 for repairing a defectin a biological tissue. The defect may include a void, an insult, and/orany other condition resulting in diminished function of the biologicaltissue. In one embodiment, method 700 includes creating 705 a void inthe biological tissue, and the defect is the created void. For example,the void may be created 705 by surgical incision to provide access to anunderlying tissue (e.g., a tumor). In another example, the void iscreated 705 by excising necrotic tissue (e.g., skin cells). One or morebiomedical patches capable of covering the defect are selected 710. Forexample, a plurality of biomedical patches may be selected 710 for alarge and/or complex (e.g., irregularly shaped) defect. The biomedicalpatch includes a plurality of radially aligned polymeric fibersextending from a center of the biomedical patch to a perimeter of thebiomedical patch. For example, a biomedical patch having a diametergreater than the diameter of an approximately circular defect may beselected 710.

The biomedical patch selected 710 may also include non-radially aligned(e.g., randomly oriented and/or uniaxially aligned) polymeric fibers.For example, radially aligned fibers and non-radially aligned fibers maybe arranged in separate layers.

In some embodiments, the biomedical patch includes multiple areas ofradially aligned fibers. In one embodiment, a first set of radiallyaligned fibers extends from a center of the biomedical patch to a firstperimeter and define an inner area. A second set of radially alignedfibers extends from the first perimeter to a second perimeter anddefines an outer area.

A substance such as a growth factor and/or a drug (e.g., achemotherapeutic drug) may be applied 715 to the biomedical patch. Forexample, the biomedical patch may be immersed in the substance to allowthe substance to occupy a cavity within hollow fibers of the biomedicalpatch, dope the polymer comprising the fibers in the biomedical patch,or coat the surface of the fibers within the biomedical patch.

The biomedical patch is applied 720 to (e.g., overlaid on) thebiological tissue to cover at least a portion of the defect. Forexample, the biomedical patch may be applied 720 to dura mater tissue,cardiac tissue, and/or any biological tissue including a defect. In oneembodiment, the perimeter of the biomedical patch extends past theperimeter of the defect, such that the entire defect is covered by thebiomedical patch. In some embodiments, the biomedical patch is coupled725 to the biological tissue with a plurality of sutures, adhesive,and/or any other means of attaching the biomedical patch to thebiological tissue. In an alternative embodiment, the biomedical patch issimply allowed to fuse to the biological tissue, such as by adhesion ofbiological cells to the biomedical patch.

After the biomedical patch is applied 720 and, optionally, coupled 725,to the biological tissue, the biological tissue is covered 730. In oneembodiment, other tissue overlaying the defect (e.g., dermis and/orepidermis) is repaired (e.g., sutured closed). In another embodiment,one or more protective layers are applied over the biological tissue.For example, a bandage may be applied to a skin graft, with or without aprotective substance, such as a gel, an ointment, and/or anantibacterial agent. In one embodiment, the protective layer includes ananofiber structure, such as an additional biomedical patch, asdescribed herein.

Embodiments described herein are operable with any neurosurgicalprocedure involving the repair, replacement, or expansion of the duramater, including, but not limited to, a transphenoidal procedure (e.g.,surgical removal of pituitary adenomas), various types of skull basesurgeries, and/or surgical removal of cranial or spinal tumors (e.g.,meningiomas and/or astrocytomas). In one embodiment, a biomedical patchmay be applied to a bone fracture (e.g., a complex fracture). In anotherembodiment, a biomedical patch may be applied to a defect in the skin(e.g. a burn).

Moreover, such embodiments are operable to provide a dura matersubstitute, a biomedical patch for a skin graft (e.g., dermal orepidermal), a biomedical patch for tracheal repair, a scaffold for anartificial heart valve leaflet, an artificial mesh for surgical repairof a gastrointestinal tract (e.g., an abdominal hernia or an ulcer), anartificial mesh for surgical repair of cardiac defects. For example, acardiac biomedical patch including radially aligned fibers may be usedto promote cardiomyocyte regeneration. Embodiments described hereinfacilitate providing a cardiac patch of sufficient flexibility to enablemovement of the biomedical patch by a biological tissue (e.g.,cardiomyocytes).

In some embodiments, a biomedical patch has a thickness less than athickness of the biological tissue being repaired. As cells migratealong the radial fibers of the biomedical patch, the biological tissueis regenerated.

Biomedical patches with radially aligned polymeric fibers facilitatereducing the expense of tissue repair, improving tissue healing time,and reducing or eliminating the risk of zoonotic infection. Moreover,such biomedical patches are relatively simple to manufacture, enablingcustomization of shape, size, and chemical composition and improvedavailability and non-immunogenicity. In addition, biomedical patcheswith radially aligned polymeric fibers exhibit excellent handlingproperties due to their cloth-like composition, eliminate the need for asecond surgery to harvest autologous graft tissue, and reduce the riskof contracture and adhesion when compared with known products.

EXPERIMENTAL RESULTS

Dura mater is a complex, fibrous membrane that consists of numerouscells and cell types, extracellular matrix proteins, and trophicfactors, all of which play an important role in the colonization andduralization of artificial dural substitutes, and the successfulimplementation of such biomedical patches in vivo. In order to evaluatethe capability of radially aligned nanofibers to interface with naturaldura, promote host cell adhesion to the graft, and enhance host cellmigration along the graft, an ex vivo model of the surgical repair of asmall dural defect was developed.

In a typical procedure, an “artificial dural defect” was introduced intoa piece of dura (1 cm×1 cm) by microsurgically cutting a small circularhole, 7 mm in diameter, in the center of the specimen. A nanofiber-basedscaffold was then utilized to repair the artificial defect by overlayingthe graft onto the dural specimen.

FIG. 13 is a schematic illustration of biological cells extending fromintact dural tissue, apposed to the edge of a scaffold, into the centralportion of the scaffold along radially-aligned nanofibers. The graftcovered the entire simulated dural defect while simultaneouslycontacting the dural tissue at the periphery of the specimen, anddemonstrates the ability of native cells in intact tissue to easilyadhere to and migrate across the nanofiber scaffolds.

FIGS. 14A-14D are a collection of fluorescence micrographs comparing themigration of cells when dural tissues were cultured on scaffolds ofradially aligned nanofibers (FIGS. 14A, 14C) and randomly orientednanofibers (FIGS. 14B, 14D) for 4 days using a custom cell culturesystem (FIG. 15). FIGS. 14C and 14D are magnified views of the centerportion shown in FIGS. 14A and 14B, respectively. The arrow marks thecenter of the scaffold.

As shown in FIG. 14A, dural fibroblasts stained with fluoresceindiacetate (FDA) migrated from the surrounding tissue along the radiallyaligned nanofibers and further to the center of the circular scaffoldafter incubation for 4 days. It was found that the cells couldcompletely cover the entire surface of the scaffold in 4 days. Incontrast, a void was observed after the same period of incubation timefor a scaffold made of random fibers (FIG. 14B), indicating fastermigration of native cells on radially-aligned nanofiber scaffolds thanthe random counterparts. It is clear that the scaffold made of radiallyaligned nanofibers (shown in FIGS. 14A and 14C) was completely populatedwith dural cells which had migrated from the borders of the apposeddural tissue. On the contrary, an acellular region is clearly visible atthe center of the scaffold made of randomly oriented nanofibers afterthe same incubation time, indicating cellular infiltration wasincomplete and occurred at a slower rate.

In order to further investigate the effect of fiber alignment andnanofiber scaffold post-modification on cell migration, primary duralfibroblasts isolated from dura tissue were cultured on scaffolds ofradially aligned and randomly oriented nanofibers with and withoutfibronectin coating. FIGS. 15A-C are schematic diagrams of a custom-madecell culture system designed to model wound healing of tissue defects.Specifically, dural fibroblasts were selectively seeded around theperiphery of a circular scaffold of nanofibers, effectively forming a7-mm “simulated dural defect” in the center of the sample.

FIGS. 16A-16D are fluorescence micrographs showing cell morphology anddistribution on scaffolds of radially aligned nanofibers (FIGS. 16A,16C) and randomly oriented nanofibers (FIGS. 16B, 16D) without and withfibronectin coating after incubation for 1 day. As shown in FIG. 16A,many cells could attach to the uncoated scaffolds including radiallyaligned nanofibers. In comparison, fewer cells poorly attached to theuncoated scaffold of randomly oriented nanofibers and cell aggregationswere noticed (FIG. 16B). Seeded cells were distributed evenly over theentire surface of the fibronectin-coated scaffold of radially alignednanofibers, and they exhibited an elongated shape parallel to the axisof nanofiber alignment (FIG. 16C). This result indicates thatfibronectin coating could enhance the influence of topographic cues oncell morphology provided by aligned fibers. The cells could also adherewell to the fibronectin-coated scaffold of randomly oriented nanofibersand cell distribution was more uniform than the uncoated samples, thoughno cell elongation or alignment was observed (FIG. 16D). The randomorganization of cells on the randomly-oriented nanofiber scaffolds alsomimics the organization of cells in scar tissue. This suggests that thealigned scaffolds may assist in reducing scar tissue formation bypromoting more regular cell organization/function.

To characterize cell motility on the scaffold, cells were stained withFDA and fluorescence images were taken at different time points. FIGS.17A-17D are fluorescence micrographs showing the migration of durafibroblasts seeded on fibronectin-coated scaffolds of radially alignednanofibers for 1 day (FIG. 17A), 3 days (FIG. 17B), and 7 days (FIG.17C). FIG. 17D is a magnified view of FIG. 17C. The cells were radiallyaligned, replicating the alignment of fibers underneath, as shown inFIG. 17D.

The ability of dural fibroblasts to migrate into and repopulate asimulated dural defect was measured at various time points throughoutthe experiment as an estimate of the regenerative capacity of thesubstitute. FIG. 18 is an illustration of the determination (e.g.,calculation) of the area of simulated dural defect remaining on thescaffold at a given time point. FIG. 19 is an illustration of the areaof void space as a function of incubation time. In FIG. 19, “Random”indicates samples with a scaffold of random fibers; “Random F” indicatessamples with a fibronectin-coated scaffold of random fibers; “Aligned”indicates samples with a scaffold of radially aligned fiber; and“Aligned F” indicates samples with a fibronectin-coated scaffold ofradially aligned fibers. An asterisk (*) and a hash (#) indicate p<0.05for samples compared with Random samples and Random F samples in thesame period of incubation time.

The area of void decreased with increasing incubation time for all thescaffolds tested due to the inward migration of cells. As illustrated byFIGS. 17A-17D, aligned fibers may significantly enhance cell migrationcompared to random fibers, and cells migrated fastest on thefibronectin-coated scaffold of radially aligned nanofibers for the first3 days of incubation. Around 5 mm² of surface area remained uncovered bycells on the uncoated random scaffolds even after incubation for 7 days.In contrast, cells covered almost the entire area of the simulateddefect within the same period of incubation for other three types ofscaffolds.

FIGS. 20A-20D are fluorescence micrographs showing live duralfibroblasts labeled with membrane dye on scaffolds of radially alignednanofibers with fibronectin coating after a 1-day culture (FIG. 20A), a3-day culture (FIG. 20B), a 7-day culture (FIG. 20C), and a 10-dayculture (FIG. 20D). FIG. 20 D includes an inset of a high magnificationimage of FIG. 20D indicating that the cells were radially aligned on thealigned scaffolds. Cell migration towards the center of afibronectin-coated scaffold of radially aligned nanofibers was furtherconfirmed by time lapse imaging shown in FIGS. 20A-20D.

Dural tissue is primarily composed of type I collagen. The production oftype I collagen from dural fibroblasts was also examined. FIGS. 21A-21Dare fluorescence micrographs obtained by immunostaining of type Icollagen with cell nuclei with 4′,6-diamidino-2-phenylindole (DAPI) inblue for scaffolds of radially aligned fibers (FIGS. 21A, 21C) andrandomly oriented fibers (FIGS. 21B, 21D). It was observed thatcomparable levels of type I collagen were produced by cells on thescaffolds of radially aligned fibers as compared to the scaffolds ofrandom fibers although one previous study showed more elongated cellsexpressed higher collagen type I than did less elongated cells.Additionally, fibronectin coating had no significant influence on theproduction of type I collagen. The type I collagen was orientedhaphazardly for the random scaffolds, resembling the extracellularcomposition of amorphous scar tissue, and had a high degree oforganization for the radially aligned scaffolds, resembling healthyconnective tissue

Recent advances in cell-biomaterial interaction have shown that bothchemical and topographical properties of the materials surface canregulate and control cell shape and function. Cell orientation,motility, adhesion and shape can be modulated by specific surface micro-and nano-topographies. Cells can align along microgrooves or similartopographical features on a surface. It was demonstrated thatfibroblasts were the most sensitive cell-type compared to endothelialcells and smooth muscle cells, and responded with a strong alignment,elongation, and migration along such topographical features.

Simultaneously, electrospinning has been widely used for producingnanofibers for a rich variety of applications in tissue engineeringincluding skin grafts, artificial blood vessels, nerve repair, andothers. Yet previous studies were limited to the use of scaffolds madeof random and uniaxially-aligned nanofibers. Scaffolds composed ofuniaxially-aligned nanofibers are not practical for wound healingapplications due to the commonality of irregularly shaped wounds. Thework described herein demonstrated for the first time the fabrication ofa new type of scaffolds made of radially aligned nanofibers. This noveltype of scaffold can guide dural fibroblasts spreading along thedirection of fiber alignment and direct cell motility towards the centerof the scaffold, resulting in faster cell migration and infiltrationcompared to scaffolds composed of randomly oriented nanofibers.

In addition, uniaxially aligned nanofiber scaffolds cannot match such acapability in that they can guide cell migration only in one direction.It was reported that controlling cellular orientation or morphology bytopography, the so-called “contact guidance”, could allow for theorganization of extracellular matrix. For most injuries, repair resultsin previously functional tissue becoming a disorganized amalgam of cell(e.g., fibroblasts) and extracellular matrix (e.g., collagen fibers)known as a scar. Highly organized cells and extracellular matrix isrequired for proper tissue regeneration and function, which is normallyvastly different from tissue repair with scarring. It has beendemonstrated in the present work that extracellular matrix type Icollagen on scaffolds of radially aligned nanofibers showed a highdegree of organization, suggesting that radially-aligned nanofiberscaffolds may reduce the possibility of scar tissue formation followingwound healing.

A dura substitute should be safe, efficacious, easy to handle,watertight, and easily integrated into the surrounding tissue to formnew tissue similar to the native tissue. Also, it should avoid harmfulforeign body reactions, be free of any potential risk of infections,have mechanical properties similar to those of natural dura mater, inparticular with respect to flexibility and strength, be stable and/orstorable, and be available for immediate use. In the present work,biodegradable polymer PCL was chosen as a material for dural substitutein that PCL has some advantages compared with other bioabsorbablepolyesters. Heterogeneous degradation of PGA and poly(L-lactic acid)(PLLA) could lead to a sudden increase of degradation products,resulting in acidic conditions and toxic reactions in the surroundingtissue. The degradation of PCL is slower and produces less-acidicdegradation products and has been studied as a wound dressing materialssince the 1970s.

In order to obtain water-tight property, the radially-aligned nanofiberscaffold can be combined with nonwoven mat to form two-layered or evenmulti-layered substitutes. Simultaneously, antibiotics can be readilyencapsulated inside nanofibers to further reduce inflammatory response,improve wound healing, and prevent postsurgery adhesion. Alternatively,PCL can be blended with other polymers to further improve itsbiocompatibility, as well as mechanical, physical, and chemicalproperties. Moreover, extracellular proteins and/or growth factors canbe immobilized on the surface of the nanofibers using various surfacemodification approaches to enhance cell adhesion. The current workdemonstrates the effect of fibronectin coating on the PCL nanofibersthrough electrostatic interaction on dural fibroblast adhesion andmotility. The results presented herein demonstrate that fibronectincoating enhanced adhesion of dural fibroblasts and improved cellmigration on randomly oriented nanofiber scaffolds. In contrast, thecoating had marginal contribution to cell motility on radially alignednanofiber scaffolds, compared to the bare scaffolds, indicating thepredominant role played by nanofiber alignment and resulting surfacetopography.

In summary, the fabrication of a new type of electrospun nanofiberscaffold including radially aligned fibers and the potential applicationof such structures as dural substitutes are described herein. Duralfibroblasts cultured on scaffolds of radially aligned nanofibers wereelongated parallel to the fiber axis, and cell migration towards thecenter of the scaffold was accelerated along with the development of aregular arrangement of extracellular matrix like type I collagen,potentially promoting fast regeneration and formation of neodura. Takentogether, these results suggest that radially aligned nanofibers possessgreat potential as an artificial dural substitute, may offer analternative in the repair of dural defects, and furthermore occupy aunique, desirable niche within the neurosurgical community.

Additional Experimental Results

In a typical procedure for electrospinning PCL (Mw=65 kDa,Sigma-Aldrich) nanofibers, a solution of 20% (w/v) PCL in a mixture ofdichloromethane (DCM) and N,N-dimethylformamide (DMF) (Fisher Chemical)with a volume ratio of 8:2 was used. The fibers were spun at 10-17 kVwith a feeding rate ranging from 0.5 mL/h, together with a 23 gaugeneedle as the spinneret. A piece of aluminum foil was used as acollector to obtain random nanofiber scaffolds. Radially alignednanofiber scaffolds were fabricated utilizing a collector consisting ofa ring electrode (e.g., metal ring) and a point electrode (e.g., a sharpneedle). Electrospun PCL nanofibers were coated with fibronectin(Millipore, Temecular, Calif.) as the following. The electrospun fiberscaffolds were sterilized by soaking in 70% ethanol overnight and washedthree times with phosphate buffered saline (PBS). Then, the scaffoldswere immersed in a 0.1% poly-_(L)-lysine (PLL) (Sigma-Aldrich) solutionfor one hour at room temperature, followed by washing with PBS buffer(Invitrogen) three times. Subsequently, the samples were immersed in afibronectin solution (26 μL 50 μg/mL fibronectin solution diluted with 5mL PBS buffer) at 4° C. overnight. Prior to cell seeding, thefibronectin solution was removed and the nanofiber scaffolds were rinsedwith PBS buffer.

The PCL nanofiber scaffolds were sputter-coated with gold before imagingwith scanning electron microscope (Nova 200 NanoLab, FEI, Oreg., USA) atan accelerating voltage of 15 kV. Samples prepared for use in cellculture were inserted into a 24-well TCPS culture plate and sterilizedby soaking scaffolds in 70% ethanol.

Fibroblasts were isolated from sections of explanted dura. Specifically,a 2.0 cm×1.5 cm section of dura was removed through sharp dissection andwashed three times with cold PBS. Dural fibroblasts were then isolatedby digesting minced dura three times in 4 mL of warm Hank's BalancedSalt Solution (HBSS) containing 0.05% Trpsin and 0.04% EDTA(Sigma-Aldrich, St. Louis, Mo.). Following digestion collectedsupernatant was centrifuged and the pellet of dural cells was isolatedand resuspended in Dulbecco's modified Eagle's medium (DMEM)supplemented with 10% calf serum and 1% penicillin and streptomycin.Dural cells obtained in this manner were then plated in 75 cm² flaks andexpanded (subpassaged no more than 5 times).

Large continuous pieces of dura mater were placed in cold PBS andmicrosurgically trimmed into 1 cm×1 cm sections. An artificial defectwas then introduced into each section of dura by microsurgically cuttinga small circular hole, 7 mm in diameter, in the middle of the section.Sections of dura were then introduced into individual wells of 6-wellculture plates containing 4 mL of DEMEM supplemented with 10% calf serumand 1% penicillin and streptomycin. Random and radially alignednanofiber scaffolds 1 cm in diameter were then utilized to repair theartificial defects by overlaying the graft onto the dural specimen.Nanofiber scaffolds were placed on the dura such that the graft coveredwith entire defect while simultaneously contacting the dural tissue atthe periphery of the specimen. Nanofiber scaffolds were held in thisposition throughout the experiment by placing a sterilized metal ringover both the scaffold and the dura. After 4 days of culture, the cellswere stained with FDA in green color and imaged with fluorescencemicroscope. Fluorescent images were taken using a QICAM Fast Cooled Mono12-bit camera (Q Imaging, Burnaby, BC, Canada) attached to an Olympusmicroscope with OCapture 2.90.1 (Olympus, Tokyo, Japan). Similarly,around 1×10⁵ dural fibroblast cells were seeded onto the periphery ofnanofiber scaffolds using the custom-made culture system shown in FIGS.15A-C. After different periods of time, the cells were stained with FDAin green color and imaged with fluorescence microscope. The totalsurface area of nanofiber scaffold devoid of migrating cells was thenquantified using Image J software (National Institute of Health).

Living cells were labeled with membrane dye using VYBRANT DiOcell-labeling solution (Invitrogen) according to the manufacturer'sinstructions and then imaged at day 1, 3, 7, and 10.

Production of collagen type I by the dural fibroblasts on the fiberscaffolds was assessed using immunohistochemistry. At day 7, the cellswere rinsed with PBS and fixed with 3.7% formalin for 1 h (N=4). Cellswere permeabilized using 0.1% Triton X-100 (Invitrogen) in PBS for 20min, followed by blocking in PBS containing 5% normal goat serum (NGS)for 30 min. Monoclonal antibodies for type I collagen (1:20 dilution)was obtained from EMD Chemicals (Calbiochem, San Diego, Calif.). Cellswere washed three times with PBS containing 2% FBS. The secondaryantibody Gt×Rb IgG Fluor (Chemicon, Temecula, Calif.) (1:200 dilution)was applied for 1 h at room temperature. Fluorescent images were takenusing a QICAM Fast Cooled Mono 12-bit camera (Q Imaging, Burnaby, BC,Canada) attached to an Olympus microscope with OCapture 2.90.1 (Olympus,Tokyo, Japan).

Mean values and standard deviation were reported. Comparative analyseswere performed using the Turkey post hoc test by analysis of variance ata 95% confidence level.

As a secondary study, an ex vivo model of the surgical repair of a smalldural defect was developed. Large pieces of healthy dura mater (3 cm×3cm) were placed into cold, supplemented Dulbecco's Modified Eagle Media(DMEM) and microsurgically trimmed into smaller (1 cm×1 cm) pieces.Artificial defects were introduced into the pieces of dura bymicrosurgically cutting small circular holes, 6-8 mm in diameter, intothe middle of the specimens. Radially aligned nanofiber scaffolds,randomly oriented nanofiber scaffolds, and DURA MATRIX collagenscaffolds (1 cm×1 cm) were then utilized to repair the artificialdefects by overlaying the graft onto the dural specimen, such that thegraft covered the entire defect while simultaneously contacting thedural tissue at the periphery of the specimen.

Assemblies of dural/dural substitute were then cultured in vitro insupplemented DMEM for a period of four days. At the terminal time point,optical and fluorescent microscopy was utilized to assess theregenerative capacity of the substitute, defined as the ability of duralcells to migrate onto the artificial substitute and repopulate theacellular region of the dural substitute within the artificial defect.

Results demonstrated that native cells present in intact dura (primarilydural fibroblasts) readily migrated onto apposed polymeric nanofiberdural substitutes in high concentrations within 24 to 48 hours aftercoming into contact with pieces of explanted dura. Dural cell migrationonto gold-standard collagen matrices followed a similar time course,though slightly lower concentrations of dural cells were observedmigrating onto collagen matrices compared to nanofiber duralsubstitutes. This observation suggests that nanofiber dural substituteseasily adhere to native dural tissue, an important quality regarding theintraoperative handling and/or placement of the material, and thatnanofiber dural substitutes provide an ideal substrate for duralfibroblast adhesion.

Further examination of the various dural substitutes after four days ofculture revealed that dural fibroblast migration into the central,acellular region of the material proceeded significantly faster onradially aligned nanofiber substitutes than on randomly orientednanofiber substitutes or collagen matrices. This finding was evidencedby the fact that after four days of culture, a prominent acellularregion (“void space”) remained on samples of both the random nanofibersubstitute and the collagen matrix.

In contrast, samples of radially aligned nanofiber materials examined atthe same time point were completely populated with dural cells which hadmigrated from the borders of the apposed dural tissue. In effect,radially aligned nanofiber substitutes were able to induce significantlyfaster “healing” of this simulated dural defect than both randomlyoriented materials. High magnification views of dural substitutes withinthis ex vivo culture further demonstrated the ability of radiallyaligned nanofiber materials to align and direct native, migratory duralcells, a result similar to that of the previous study conducted usingpre-seeded dural fibroblasts. Specifically, dural cells were noted toalign and extend parallel to individual nanofibers within the artificialsubstrate, as well as deposit organized extracellular matrix proteins(namely type I collagen) on the aligned nanofiber materials. Thisobservation suggests that the topographical cues presented by alignednanofiber substitutes are capable of organizing and directing nativedural cells migrating from intact dura, and may enhance the ability ofthese migratory cells to deposit extracellular matrix proteins necessaryto heal and repair dural defects.

Results of this secondary study demonstrate that nanofiber duralsubstitutes not only provide a favorable scaffold for dural celladhesion and migration, but readily support the ingrowth of dural cellsfrom whole, intact dura mater. The ability of nanofiber materials tointimately interface intact dura mater and facilitate rapid cellularpopulation of the polymeric scaffold strongly suggest that this materialmay function exceptionally well as an artificial graft in the surgicalrepair of complex dural defects. In addition, dural substitutesconstructed of radially aligned nanofibers were demonstrated to promotefaster “healing” of simulated dural defects than randomly orientedmaterials, suggesting that aligned nanofiber scaffolds impartingnanoscale topographical features may represent a significanttechnological advance over clinical gold-standard collagen matrices.

Although experiments described herein were limited in duration, theresults of these experiments suggest that biomedical patches includingradially aligned fibers are viable for use in tissue repair at longerdurations. For example, it is expected that the observed acceleratedcellular ingrowth would continue until the biological tissue at the siteof a defect is completely regenerated and/or until degradation of thebiomedical patch is complete.

In Vivo Experimental Results

In vivo experimentation was performed by imposing a 12 millimeterdiameter dural defect in native porcine dura. The defect was repairedwith a collagen dural substitute, a mono-layer dural substitute withrandomly oriented nanofibers, and a bi-layer dural substitute with onelayer of radially aligned nanofibers fused to a second layer of randomlyoriented nanofibers through layer-by-layer stacking (e.g., as describedabove with reference to FIG. 8). In a control group, the defect wasunrepaired.

FIG. 22 is a graph 2200 illustrating the thickness of regenerated duraat the center of repaired dural defects over time. In graph 2200, ay-axis 2205 represents the total thickness of regenerated dura,including both regenerative tissue and the integrated dural substitutematerial, at the center of a dural defect. Samples with no duralsubstitute (control samples), a collagen dural substitute, a mono-layerrandomly oriented nanofiber dural substitute, and a bi-layer radiallyaligned nanofiber dural substitute are grouped by elapsed time on anx-axis 2210.

FIG. 23 is a graph 2300 illustrating regenerative collagenous tissuecontent over time. In graph 2300, a y-axis 2305 represents thepercentage of regenerated dura that is composed of regenerativecollagenous tissue. Samples with a collagen dural substitute, amono-layer randomly oriented nanofiber dural substitute, and a bi-layerradially aligned nanofiber dural substitute are grouped by elapsed timeon an x-axis 2310.

Electrode Arrays

In some embodiments, a collector includes a plurality of electrodes atleast partially circumscribing an area and a second electrode positionedwithin the area. The electrodes may be arranged in an array, such as agrid and/or other polygonal pattern, and a polymer deposited on theelectrodes may form fibers extending between the electrodes of thecollector, such that the fibers define the sides of a plurality ofpolygons, with the electrodes positioned at the vertices of thepolygons. In some embodiments, the structure created by such fibers maybe used to create a cell microarray, such as by seeding the structurewith cells and incubating the cells to promote propagation of the cellsthroughout the structure.

Cell microarrays may provide powerful experimental tools forhigh-throughput screening useful in a number of applications rangingfrom drug discovery and toxicology to stem cell research and tissueengineering. For example, cell microarrays may represent an effectivemeans of fabricating ordered neuronal networks useful in studyingsynapse formation and neuronal plasticity in vitro. At least some knowntechniques for fabrication of neuronal microarrays have concentrated onthe use of spatial patterning of cell adhesive and/or cell repulsivematerials and agents. Unfortunately, such fabrication techniques may betime consuming and costly, and involve the use of sophisticatedinstrumentation (e.g., photolithography, soft lithography, contactprinting, microfluidics, nanoprinting, and inkjet printing).

Electrospinning is capable of producing one-dimensional fibers withdiameters ranging from several nanometers to several microns. The largesurface area to volume ratio and nanoscale morphology of electrospunnanofibers may suggest that these materials effectively mimic thearchitecture of extracellular matrix (ECM). As a result, electrospunnanofiber materials have been utilized in a wide variety of biomedicalapplications. Electrospun nanofibers may be deposited on a conductivecollector in a random fashion and/or aligned into uniaxial arraysthrough manipulation of an electric field and/or application ofmechanical force.

Embodiments described herein facilitate producing a complex cellmicroarray using electrospun nanofibers. In exemplary embodiments, acollector with an array of electrodes is used to fabricate electrospunnanofiber scaffolds that include a complex, ordered architecture andnumerous multiwells. Such a scaffold may be valuable at least for i)cell microarray formation; and ii) neuronal network formation. The useof presented complex nanofiber arrays may facilitate the creation ofadvanced substrates useful in neural engineering applications and cellarrays useful in bio-sensing and drug screening applications.

FIG. 24 is a diagram illustrating a perspective view of an exampleelectrospinning system 2400 for producing a structure of polygonallyaligned fibers using an array of electrodes. System 2400 is similar tosystem 100 (shown in FIG. 1) in structure and operation. A collector2405 includes a plurality of first electrodes 2410, which may bereferred to as peripheral electrodes. First electrodes 2410 defineand/or at least partially circumscribe an area 2415, such as a polygon.As illustrated in FIG. 24, the area 2415 defined by first electrodes2410 is a hexagon. A second electrode 2420, which may be referred to asan inner electrode, is positioned within (e.g., approximately at thecenter of) area 2415, such that first electrodes 2410 surround secondelectrode 2420. In exemplary embodiments, first electrodes 2410 andsecond electrodes are metallic (e.g., stainless steel) beads having adiameter between 0.5 millimeters (mm) and 5.0 mm (e.g., 1.0 mm or 2.0mm).

System 2400 also includes a spinneret 120 and is configured to create anelectric potential between collector 2405 and spinneret 120, asdescribed above with reference to FIG. 1. In exemplary embodiments,peripheral electrodes 2410 and inner electrode 2420 are electricallycoupled to a power supply 130 via a conductor 135, and spinneret 120 iscoupled to power supply 130 via a conductor 145. Power supply 130 isconfigured to charge peripheral electrodes 2410 at a first amplitudeand/or polarity via conductor 135, and to charge spinneret 120 at asecond amplitude and/or polarity, opposite the first polarity, viaconductor 145.

In the embodiment illustrated in FIG. 24, peripheral electrodes 2410 andinner electrode 2420 are metallic (e.g., stainless steel) beads orballs, which may be referred to as “microbeads,” arranged in a hexagonalpattern. In some embodiments, circular enclosed area 125 may have adiameter of between 1 centimeter and 20 centimeters. In otherembodiments, peripheral electrodes 2410 and inner electrode 2420 may beany shape and/or may be arranged in any pattern suitable for use withthe methods described herein. For example, peripheral electrodes 2410and inner electrode 2420 may be pins, rods, domes, and/or ridges.Further, peripheral electrodes 2410 and inner electrode 2420 may bearranged in an octagonal, pentagonal, and/or square pattern, forexample, though other polygonal and non-polygonal arrangements, regularand/or irregular, are also contemplated.

In one embodiment, area 2415 defines a horizontal plane 2425. Spinneret120 is aligned with inner electrode 2420 and vertically offset fromhorizontal plane 2425 at a variable distance. For example, spinneret 120may be vertically offset from horizontal plane 2425 at a distance of 1centimeter to 100 centimeters. In exemplary embodiments, inner electrode2420 and/or peripheral electrodes 2410 include a rounded (e.g., convex)surface, such as the surface of the metallic beads shown in FIG. 24,oriented toward horizontal plane 2425.

As described above with reference to FIG. 1, spinneret 120 is configuredto dispense a polymer 140 while spinneret 120 is electrically charged atthe second amplitude and/or polarity, and peripheral electrodes 2410 andinner electrode 2420 are electrically charged at the first amplitudeand/or polarity. Spinneret 120 dispenses polymer 140 as a stream 160.Stream 160 has a diameter approximately equal to the aperture diameterof spinneret 120. Stream 160 descends toward collector 2405. Forexample, stream 160 may fall downward under the influence of gravityand/or may be attracted downward by a charged conductive surface 162positioned below collector 2405. For example, conductive surface 162 maybe electrically coupled to conductor 135 and charged at the sameamplitude and/or polarity as peripheral electrodes 2410 and centralelectrode 2420. As stream 160 descends and is deposited on collector2405, polymer 140 forms one or more solid polymeric fibers 2430extending from inner electrode 2420 to a peripheral electrode 2410and/or between peripheral electrodes 2410.

In some embodiments, collector 2405 includes peripheral electrodes 2410that define a plurality of areas 2415. For example, peripheralelectrodes 2410 immediately surrounding inner electrode 2420 may beconsidered inner peripheral electrodes, and a plurality of outerperipheral electrodes 2435 may surround inner peripheral electrodes2410, such that inner peripheral electrodes 2410 are nested within outerperipheral electrodes 2435. Collector 2405 may include any quantity ofnested sets of peripheral electrodes. While collector 2405 includeselectrodes in a closely-packed arrangement (e.g., with electrodescontacting each other), it is contemplated that electrodes may bedisplaced from each other by an inter-electrode distance, which may beconstant throughout the collector or may vary between different pairs ofelectrodes.

Further, in some embodiments, a collector may include electrodes thatdefine a plurality of partially overlapping areas in a modular fashion.FIG. 25 is a diagram illustrating a perspective view of an examplemodular electrospinning collector 2500. Collector 2500 includes firstelectrodes 2505 surrounding a second electrode 2510. First electrodes2505 define a first hexagonal area 2515. With respect to first hexagonalarea 2515, second electrode 2510 may be considered an inner electrode,and first electrodes 2505 may be considered peripheral electrodes.

Collector 2500 also includes a plurality of third electrodes 2520 thatare positioned outside first hexagonal area 2515. Third electrodes 2520,second electrode 2510, and a subset of first electrodes 2505 define asecond hexagonal area 2525 that partially overlaps first hexagonal area2515. One of the first electrodes 2505 (e.g., a peripheral electrodewith respect to first hexagonal area 2515) is positioned within secondhexagonal area 2525. With respect to second hexagonal area 2525, thisfirst electrode 2505 may be considered an inner electrode. Thirdelectrodes 2520, the subset of the first electrodes 2505, and the secondelectrode 2510 may be considered peripheral electrodes. Althoughelectrodes defining two partially overlapping areas are illustrated inFIG. 25, it is contemplated that the modular nature of collector 2500facilitates including any quantity of electrodes that define anyquantity of areas, such that collector 2500 may be extended in one ormore directions by adding electrodes to the perimeter of collector 2500.

As described above with reference to system 2400 (shown in FIG. 24),collector 2500 (e.g., first electrodes 2505, second electrode 2510, andthird electrodes 2520) is configured to be electrically charged at anamplitude and/or a polarity opposed the amplitude and/or polarity atwhich spinneret 120 is electrically charged. When these components areso charged, a polymer dispensed by spinneret 120 may form fibersextending between the electrodes (e.g., first electrodes 2505, secondelectrode 2510, and/or third electrodes 2520) of collector 2500.

FIG. 26 is a diagram 2600 illustrating an electric field generated by anelectrospinning system such as electrospinning system 2400 (shown inFIG. 24). Diagram 2600 shows a two dimensional, cross-sectional view ofelectric field strength vectors between a spinneret 120 and a pluralityof electrodes 2605.

Electric field vectors near the surface of electrodes 2605 are orientedperpendicular to the surface of electrodes 2605. Electric field vectorsbetween two neighboring electrodes split into two main streams, pointingtowards the centers of the two adjacent electrodes 2605. Accordingly,fibers deposited on the surface of electrodes 2605 may be randomlydistributed, while the fibers deposited in the region between twoneighboring electrodes 2605 may be uniaxially aligned between these twoadjacent electrodes 2605.

FIGS. 27A-27F are microscopy images of a nanofiber membrane 2705produced using a collector with an array of electrodes, such ascollector 2405 (shown in FIG. 24). For example, membrane 2705 may beproduced using an array of stainless steel beads. FIG. 27A is an opticalmicroscopy image of a membrane 2705. FIG. 27A includes an inset 2710illustrating a magnification of membrane 2705 with a light source on theright-hand side of the image. Shadows in inset 2710 indicate wellswithin membrane 2705, the positions of which correspond to the positionsof electrodes in the collector.

FIG. 27B is a scanning electron microscopy (SEM) image of membrane 2705illustrating the complex, ordered architecture composed of hexagonallyarranged wells 2715 connected with uniaxially aligned fiber arrays 2720.The depth of the wells formed by depositing electrospun nanofibers onpacked stainless steel microbeads 1.0 mm and 2.0 mm in diameter wasapproximately 200 micrometers (μm) and 400 μm, respectively. Such wellsmay be referred to as “microwells.”

FIGS. 27C-27F are magnifications of corresponding areas within FIG. 27B.FIG. 27C suggests that the fibers deposited on the surface of microbeadelectrodes were randomly distributed. FIG. 27D shows that the fibers atthe interface between the surface of an electrode and a gap betweenelectrodes transitioned from a random orientation to an alignedorientation. FIG. 27E indicates that fibers deposited along the axisconnecting the centers of two adjacent electrodes were uniaxiallyaligned parallel to that axis. FIG. 27F shows that the fiber density wassignificantly lower between neighboring beads and away from the axesconnecting the centers of adjacent beads than in other regions (e.g.,shown in FIGS. 27C-27E), and that fiber deposited in this region wererandomly oriented.

In some embodiments, a fiber membrane, such as membrane 2705, may becombined with other membranes. For example, a membrane with a pluralityof wells interconnected by uniaxially aligned fibers may be used as onelayer within a multi-layer structure, as described above with referenceto FIG. 8. In addition, or alternatively, different collector types maybe combined, such as by using an electrode array collector as an innercollector (e.g., corresponding to a center of a biomedical patch, andusing a ring-type collector (e.g., as shown in FIG. 1) as an outercollector that surrounds the inner collector.

Experimental Results

Fiber membranes, or “scaffolds,” produced by an electrode arraycollector as described above were evaluated for use as substrates forgenerating cell microarrays. Cells were selectively seeded onto thesurface of the scaffold by placing a small amount of media, containingspecified number of cells, onto the microwells present within thenanofiber arrays.

FIGS. 28A-28D are microscopy images illustrating cell growth in amembrane such as membrane 2705 (shown in FIGS. 27A-27F). FIG. 28A is anoptical microscopy image illustrating that droplets 2805 containingcells may be placed within the wells of a fiber membrane. Further,hydrophobic fibers may facilitate maintaining such droplets for over twohours. Cells adherent to the nanofiber matrices after two hours werefound to be loosely attached and were easily removed using PBS buffer,suggesting fast, reversible binding of cells within the microarrays.Cells adherent to the nanofiber matrices after twenty-four hours werestained with fluorescein diacetate (FDA) in green to identify livingcells.

FIGS. 28B-28D are fluorescence microscopy images illustrating cellmicroarrays. Live MG-63 cells were stained with fluorescein diacetateand are shown as light areas against a dark background in FIGS. 28B-28D.

FIG. 28B shows an array of cells selectively adhered to the microwellswithin the nanofiber membrane. Each well within the scaffold wasobserved to contain approximately 45 cells, while very few cells wereobserved outside of the microwells within the fiber membrane. Theaverage number of cells adherent on each microwell was easilymanipulated by controlling the density of cells present within theseeding droplets.

FIG. 28C demonstrates cell microarrays seeded with greater numbers ofcells (approximately 150 cells per well) than were used in the arraysshown in FIG. 28B. Despite increasing cell concentrations, cellsremained greatly confined to the wells in the nanofiber scaffold. FIG.28D shows the same cell microarray shown in FIG. 28C after incubationfor three days. Comparison of FIG. 28D to FIG. 28C demonstrates thatseeded cells were capable of proliferating and migrating on the surfaceof the nanofiber scaffolds, yet generally remained physically confinedwithin the wells of the cell microarray.

In order to examine the potential of these unique nanofiber scaffolds aseffective substrates for neural engineering applications, dorsal rootganglia (DRG) were seeded onto fiber membranes functionalized withpolylysine and laminin and incubated for 6 days. Resulting neuritefields protruding from DRG were stained with anti-neurofilament 200 tovisualize neurite extension along the underlying nanofiber scaffold.

FIGS. 29A and 29B are microscopy images illustrating neurite propagationin a membrane such as membrane 2705 (shown in FIGS. 27A-27F). FIG. 29Ais an overlay of an optical microscopy image and a fluorescencemicroscopy image illustrating that neurites emanated from a DRG mainbody located at the center of FIG. 29A and formed an appreciableneuronal network after 6 days of culture. Neurites were observed to growalong the long axes of uniaxially aligned nanofibers and reachneighboring microwells, effectively replicating the geometry of theunderlying nanofiber architecture.

FIG. 29B is an overlay of an optical microscopy image and a fluorescencemicroscopy image adjacent to the region shown in FIG. 29A. FIG. 29Bdemonstrates that neurites may continue growing along the direction ofuniaxial alignment of nanofibers after reaching the neighboring wellsand navigate to other neighboring wells along the fiber alignment inseveral directions. Neurites extending to adjacent microwells weresubsequently observed to split into five groups following the alignedfiber arrays which connected to a secondary set of adjacent wells,further indicating capability of the scaffold to form a complex neuronalnetwork in vitro.

FIGS. 30A and 30B are overlays of an optical microscopy image and afluorescent microscopy image illustrating neuronal network formationfrom embryoid bodies in a membrane such as the membrane shown in FIGS.27A-27F. Embryonic stem (ES) cells, cultured to aggregate into embryoidbodies (EBs) using the 4−/4+ protocol, were seeded onto electrospunnanofiber scaffolds such as that shown in FIGS. 27A-27F, and incubatedwith B27 supplement to induce neuronal differentiation. Immunostainingwith Tuj1, a neuronal marker, was performed after incubation for 14 daysto examine the ability of underlying nanofiber scaffolds to promoteneuronal differentiation in vitro.

FIGS. 30A and 30B demonstrate the ability of EBs to form neuronalnetworks on nanofiber membrane substrates. In one case, one EB wasconfined within one of the microwells, while neurites extendedperipherally along the underlying fiber pattern, as shown in FIG. 30A.Neurites extending from cultured EBs were similarly aligned on theuniaxial portion of the scaffold where fibers were highly organized.Upon reaching the region of the adjacent wells, neurites werehaphazardly organized as a result of the random orientation of theunderlying fibers.

In another case, EBs were seeded on regions of uniaxially alignednanofibers within the nanofiber array, as shown in FIG. 30B. Neuritesagain extended along the direction of fiber alignment, and, uponreaching the nearest well, exhibited a disordered organization. Notably,when the neurites extended through the microwell region, their uniaxialalignment, parallel to the underlying fiber alignment, was restored.Together, these results suggest that nanofiber architectures describedherein represent a simple and effective means of developing complexneuronal networks from either primary neurons or embryonic stem cells.

Experimental Procedure

The electrospinning system used for fabricating and collecting alignednanofibers was similar to system 2400 (shown in FIG. 24). The polymersolution used for electrospinning contained 20% PCL (w/v) in a mixedsolvent of dichloromethane (DCM) and dimethylformaldehyde (DMF) with avolume ratio of 80:20. The collector included assemblies of stainlesssteel microbeads with diameters of 1 mm and 2 mm, respectively. Thefiber membranes were transferred to culture plates and then fixed bymedical grade silicon adhesive. The PCL fibers were sputter-coated withgold before imaging with scanning electron microscope at an acceleratingvoltage of 15 kV.

For dorsal root ganglia (DRG) culture and immunostaining, DRG weredissected from the thoracic region of embryonic day 8 chicks (E8, stageHH35-36) and collected in Hank's buffered salt solution (HBSS) prior toplating. DRG were seeded on the fiber architectures and incubated for 6days in modified neurobasal (NB) media containing 1% ABAM, 1% N-2supplement (Invitrogen), and 30 ng/mL Beta nerve growth factors (B-NGF)(R&D Systems, Minneapolis, Minn.). After incubation for 6 days, the DRGwere immunostained with the marker anti-neurofilament 200(Sigma-Aldrich). Briefly, the DRG were fixed in 3.7% formaldehyde for 45minutes and permeabilized by 0.1% Triton X-100 for 30 minutes. Thesamples were blocked in PBS containing 2.5% bovine serum albumin (BSA)(Sigma-Aldrich) for 1 hour. Anti-NF 200 diluted with PBS containing 1.5%BSA was applied to the cells overnight at 4° C. A secondary antibody,AlexaFluor 488 goat anti-mouse IgG (1:200, Invitrogen), was then appliedfor 1 hour at room temperature. After staining, fluorescence images werecaptured.

For embryoid body formation and immunostaining, EBs were seeded ontofiber architectures and incubated with neural basal media containing B27supplement. After 14 days, immunohistochemistry was performed tovisualize the spatial distribution of neurites according to our previousstudy.

The MG-63 cell line was used to demonstrate the formation of cellmicroarrays. Cells were cultured in alpha minimum essential medium(α-MEM, Invitrogen, Grand Island, N.Y.), supplemented with 10% fetalbovine serum (FBS, Invitrogen) and 1% antibiotics (containing penicillinand streptomycin, Invitrogen). The medium was changed every other day,and the cultures were incubated at 37° C. in a humidified atmospherecontaining 5% CO₂. A certain number of cells were seeded into each wellof the scaffolds by placing small droplets onto wells. After incubationfor 2 hours, the scaffolds were washed with culture media to remove theloosely attached cells. Then, the living cells were stained withfluorescein diacetate (FDA) after incubation for 24 hours and imagedwith fluorescence microscope.

Additional Electrode Array Arrangements

In addition to particular examples of electrode arrays described abovewith reference to experimental results, it is contemplated thatnanofiber structures such as those described herein may be produced withvarious other electrode arrays. FIGS. 31A-31D are scanning electronmicroscopy images illustrating membranes produced using a variety ofelectrode arrays.

FIG. 31A illustrates a fiber membrane fabricated using a collectorcomposed of hexagonal arrays of stainless steel beads. FIG. 31Billustrates a fiber membrane fabricated using a collector composed ofhexagonal arrays of stainless steel beads having a larger diameter thanthe stainless steel beads used to produce the membrane shown in FIG.31A.

Other, non-hexagonal, packing orders may also be employed with theelectrodes to achieve different geometries. FIG. 31C shows a fibermembrane fabricated using a collector composed of a close-packed squarearray of stainless steel beads. FIG. 31D shows a fiber membrane producedusing a collector composed of square arrays of stainless steelmicrobeads with a gradual increase of the inter-electrode distance inone direction. The fiber membranes were not removed from the collectorsduring SEM imaging and can be readily removed (e.g., peeled off) fromcollectors as needed.

FIG. 32 is a diagram of a collector 3200 with peripheral electrodes 3205partially circumscribing an area 3210. Collector 3200 also includes aninner electrode 3215. Peripheral electrodes 3205 and inner electrode3215 define a portion 3220 of area 3210. In exemplary embodiments,peripheral electrodes 3205 are positioned on a perimeter 3225 of area3210.

In the embodiment shown in FIG. 32, area 3210 is shown as an ellipse(e.g., a circle), and portion 3220 is shown as a sector of the ellipse.It is contemplated that area 3210 may be any geometric or non-geometricshape, such as an ellipse, polygon, oval, rectangle, square, triangle,and/or any rectilinear or curvilinear shape, and that portion 3220 maybe any portion of such a shape.

Electrode array fiber structures described herein enable the formationof “dimple” structures within a fiber membrane. Accordingly, theproduction of such membranes represents a significant advance in thatthe fiber membranes described possess multiple microwells arranged intovariable, ordered geometries. Furthermore, such structures possessunique, three-dimensional microwells capable of physically confiningcells seeded on the surface of the scaffold and facilitating thefabrication of cell microarrays. Compared to known approaches tomicroarray fabrication, the use of fiber membranes may be a simpler andless expensive technique for forming complex cell microarrays for invitro and in vivo use. Further, experimental results described abovedemonstrate that the neurites on the site of wells presented randomdistribution, and that neurites could bridge from one well to anotheralong the aligned fibers in between. A neuronal network developed usingsuch a structure could be used for high-throughput applications inneurotoxicology and neurodevelopmental biology.

While the making and use of various embodiments of the invention arediscussed in detail above, the embodiments of the invention provide manyapplicable inventive concepts that may be embodied in a wide variety ofspecific contexts. The specific embodiments discussed herein are merelyillustrative of specific ways to make and use the invention and do notdelimit the scope of the invention.

To facilitate the understanding of this invention, a number of terms aredefined below. Terms defined herein have meanings as commonly understoodby a person of ordinary skill in the areas relevant to the embodimentsof the invention. Terms such as “a,” “an” and “the” are not intended torefer to only a singular entity, but include the general class of whicha specific example may be used for illustration. The terminology hereinis used to describe specific embodiments of the invention, but theirusage does not delimit the invention, except as outlined in the claims.

The order of execution or performance of the operations in embodimentsof the invention illustrated and described herein is not essential,unless otherwise specified. For example, it is contemplated thatexecuting or performing a particular operation before, contemporaneouslywith, or after another operation is within the scope of aspects of theinvention. Embodiments of the invention may include additional or feweroperations than those disclosed herein.

What is claimed is:
 1. A three-dimensional electrospun nanofiberscaffold for use in repairing a defect in a tissue substrate, thethree-dimensional electrospun nanofiber scaffold comprising: a flexibledeposited electrospun fiber network of varying density, the depositedelectrospun fiber network comprising: a first set of depositedelectrospun fibers comprising a first bioresorbable polymer, wherein thefirst bioresorbable polymer comprises polyglycolic acid; and a secondset of deposited electrospun fibers comprising a second bioresorbablepolymer, the second set of deposited electrospun fibers coupled to thefirst set of deposited electrospun fibers, wherein the firstbioresorbable polymer comprises a different composition from the secondbioresorbable polymer, wherein a first portion of the flexible depositedelectrospun fiber network comprises a higher density of fibers than asecond portion of the flexible deposited electrospun fiber network, andwherein the first portion comprises a higher tensile strength than thesecond portion; wherein the three-dimensional electrospun nanofiberscaffold comprises varying density to be sufficiently flexible tofacilitate application of the three-dimensional electrospun nanofiberscaffold to uneven surfaces of the tissue substrate; wherein thethree-dimensional electrospun nanofiber scaffold comprises varyingdensity to be sufficiently flexible to enable movement of thethree-dimensional electrospun nanofiber scaffold by the tissuesubstrate, wherein the first set of deposited electrospun fibers and thesecond set of deposited electrospun fibers are configured to degradewithin three months after application to the tissue substrate, andwherein each fiber of the deposited electrospun fiber network comprisesa diameter of 1-1000 nanometers.
 2. The three-dimensional electrospunnanofiber scaffold of claim 1, wherein the second bioresorbable polymercomprises caprolactone.
 3. The three-dimensional electrospun nanofiberscaffold of claim 1, wherein at least one of the first set of depositedelectrospun fibers and the second set of deposited electrospun fibersare radially aligned.
 4. The three-dimensional electrospun nanofiberscaffold of claim 1, wherein at least one of the first set of depositedelectrospun fibers and the second set of deposited electrospun fibersare non-radially aligned.
 5. The three-dimensional electrospun nanofiberscaffold of claim 1, wherein at least one of the first set of depositedelectrospun fibers and the second set of deposited electrospun fibersare randomly oriented.
 6. The three-dimensional electrospun nanofiberscaffold of claim 1, wherein the deposited electrospun fiber networkcomprises a single layer.
 7. The three-dimensional electrospun nanofiberscaffold of claim 1, wherein the deposited electrospun fiber networkcomprises multiple layers.
 8. A three-dimensional electrospun nanofiberscaffold for use in repairing a defect in a tissue substrate, thethree-dimensional electrospun nanofiber scaffold comprising: a flexibledeposited electrospun fiber network of varying density, the depositedelectrospun fiber network comprising: a first set of depositedelectrospun fibers comprising a first bioresorbable polymer, wherein thefirst bioresorbable polymer comprises glycolic acid; and a second set ofdeposited electrospun fibers comprising a second bioresorbable polymer,the second set of deposited electrospun fibers coupled to the first setof deposited electrospun fibers, wherein the first bioresorbable polymercomprises a different composition from the second bioresorbable polymer,wherein at least a first portion of the flexible deposited electrospunfiber network comprises a higher density of fibers than a second portionof the flexible deposited electrospun fiber network, and wherein thefirst portion comprises a higher tensile strength than the secondportion; wherein the three-dimensional electrospun nanofiber scaffoldcomprises varying density to be conformable to the defect in the tissuesubstrate, wherein the three-dimensional electrospun nanofiber scaffoldcomprises varying density to be sufficiently flexible to enable movementof the three-dimensional electrospun nanofiber scaffold by the tissuesubstrate, wherein the first set of deposited electrospun fibers and thesecond set of deposited electrospun fibers are configured to separatefrom each other within three months after application to the tissuesubstrate, and wherein each fiber of the deposited electrospun fibernetwork comprises a diameter of 1-1000 nanometers.
 9. Thethree-dimensional electrospun nanofiber scaffold of claim 8, wherein thesecond bioresorbable polymer comprises caprolactone.
 10. Thethree-dimensional electrospun nanofiber scaffold of claim 8, wherein atleast one of the first set of deposited electrospun fibers and thesecond set of deposited electrospun fibers are radially aligned.
 11. Thethree-dimensional electrospun nanofiber scaffold of claim 8, wherein atleast one of the first set of deposited electrospun fibers and thesecond set of deposited electrospun fibers are non-radially aligned. 12.The three-dimensional electrospun nanofiber scaffold of claim 8, whereinat least one of the first set of deposited electrospun fibers and thesecond set of deposited electrospun fibers are randomly oriented. 13.The three-dimensional electrospun nanofiber scaffold of claim 8, whereinthe deposited electrospun fiber network comprises a single layer. 14.The three-dimensional electrospun nanofiber scaffold of claim 8, whereinthe deposited electrospun fiber network comprises multiple layers.
 15. Amonolayer electrospun nanofiber patch for use in repairing a defect in atissue substrate, the monolayer electrospun nanofiber patch comprising:a flexible deposited electrospun fiber network of varying density, thedeposited electrospun fiber network comprising: a first set of depositedelectrospun fibers comprising a first bioresorbable polymer, wherein thefirst bioresorbable polymer comprises glycolic acid; and a second set ofdeposited electrospun fibers comprising a second bioresorbable polymer,the second set of deposited electrospun fibers coupled to the first setof deposited electrospun fibers, wherein the first bioresorbable polymercomprises a different composition from the second bioresorbable polymer,wherein an inner portion of the monolayer electrospun nanofiber patchcomprises a different fiber density than an outer portion of themonolayer electrospun nanofiber patch; wherein the monolayer electrospunnanofiber patch comprises varying density to be conformable to thedefect in the tissue substrate, wherein the monolayer electrospunnanofiber patch comprises varying density to be sufficiently flexible toenable movement of the monolayer electrospun nanofiber patch by thetissue substrate, wherein the first set of deposited electrospun fibersand the second set of deposited electrospun fibers are configured todegrade within three months after application to the tissue substrate,and wherein each fiber of the deposited electrospun fiber networkcomprises a diameter of 1-1000 nanometers.
 16. The monolayer electrospunnanofiber patch of claim 15, wherein the second bioresorbable polymercomprises caprolactone.
 17. The monolayer electrospun nanofiber patch ofclaim 15, wherein at least one of the first set of deposited electrospunfibers and the second set of deposited electrospun fibers are radiallyaligned.
 18. The monolayer electrospun nanofiber patch of claim 15,wherein at least one of the first set of deposited electrospun fibersand the second set of deposited electrospun fibers are non-radiallyaligned.
 19. The monolayer electrospun nanofiber patch of claim 15,wherein at least one of the first set of deposited electrospun fibersand the second set of deposited electrospun fibers are randomlyoriented.
 20. The monolayer electrospun nanofiber patch of claim 15,wherein the first set of deposited electrospun fibers and the second setof deposited electrospun fibers are non-radially aligned.